- Data Analysis
- Other Protocols
Quantification of Western Blots Using ImageJ
NIH ImageJ is probably the cheapest and easiest way to quantify western blots. Images should be scanned in grayscale at a resolution of a least 600 dpi. I usually scan them as tif files at 600 dpi and have never had a problem. Underexposed images will work just fine at this resolution for quantification.
Download and install ImageJ software according to the instructions on the NIH website. Once you have it installed open the software and select the image file you want to quantify. For this tutorial, I’m going to use an illustrated western blot for quantification, but as long as you stick the the underexposed blots, any western can be quantified.
As a side-note, the reason I’m using an illustration for this tutorial is because my research is funded by the federal government, and I don’t want to post one of my westerns on a website with advertisements. Feel free to contact me directly and I can give you a list of publications in which I’ve performed this western blot quantification.
For the tutorial, I’ll use the instructions “File → Open” to indicate menu and sub-menu selections that you should make.
To begin, select the rectangular selection tool on the left side of the tool menu.
Next, draw your rectangle around the band in your first lane. Once you have your band in a rectangle select Analyze → Gels → Select First Lane (or “control-1″ in Windows). This will draw a box around the band.
Keep in mind the following points:
- The software makes the top of the boxes even for every lane. So make sure the bands are horizontal. If the bands are slanted, use the Image → Rotate → Arbitrarily… commands to make them level.
- The width of the selection box is fixed for the entire analysis after selecting the first lane. So make sure the rectangle is wide enough to accommodate the widest bands.
After selecting the first lane, take the cursor and move it within the box set in the first lane. This will allow you to click-and-hold, and drag the box to the second lane (Figure below, left). Then, select the Analyze → Gels → Select Next Lane command (“control-2″ in Windows). This will select an area in the second lane that is exactly equal to the area selected in the first lane.
Repeat the previous step to select the band in the third lane. Again, while the rectangle tool is still selected, click within the yellow box and drag the box so that it covers the next band. Once this is done, use the Analyze → Gels → Select Next Lane command to select it as the third lane.
For this tutorial, we’ll assume that there are only three lanes. However, you can repeat the lane selection process for all of your western blot lanes.
After the last lane, you need to select the Analyze → Gels → Plot Lanes command. A window with will pop open. This window is the densitometry measurements. It gives a graphical depiction of the average intensity of pixels from the top of the rectangle to the bottom of the rectangle (left to right on the plot).
To measure the density of your western blot band, you want to measure the area under the peak from this plot. To do so, select the straight line tool from the main menu tools.
With the straight line tool selected, go back to the window with your lane plots. Use the straight line tool to mark off the area under the peak for the first lane. You’ll want to close off all of the peak area that rises above the background level.
After selecting the first peak, scroll down within the plot window and repeat the process to close off the remaining peaks. Once you have done selected peaks for every lane, go back to the main menu and select the wand tool.
After selecting the wand tool, go back to the window with the plots of lane peaks. Click inside each peak, starting from the top, with the wand tool. You will see the peak become outlined in yellow. This integrates the area under the curve for each peak. As you continue to select the peaks, another window labeled “Results” will open.
This window contains all of the values for the areas under the curves for each selection you make. These values can then be used to make relative comparisons of band intensity between lanes. This comparison is an indication of the relative amounts of protein that were loaded into each well.
To calculate the relative values, simply divide each value by the lane that you want to be the standard. For example, in the gel quantified in this tutorial, the first lane is our relative comparison.
Relative levels are calculated as:
- 2484.426 / 2484.426 = 1
- 4422.426 / 2484.426 = 1.78
- 7785.376 / 2484.426 = 3.13
So, using our first lane as a relative comparison, the second lane contains approximately 75% more protein than the first lane, and the third lane contains approximately 213% more protein than the first lane.
And that’s it! If you’ve followed through until this point, you’ve just completed your first densitometric analysis. Densitometry is extremely important, as it allows us to make relative comparisons and average values for the relative abundance of proteins between various samples.
Remember to always use a loading control to normalize your values. Proteins like GAPDH, tubulin, and actin are good loading controls because their levels rarely change in cells. When you use those as loading controls, divide the peak values of your target protein by the peak values of your loading control before doing a relative comparison.
If you have any further questions about the use of ImageJ to quantify western blots or densitometry in general, please feel free to leave a comment or question on this page.