DENSITOMETRY OF WESTERN BLOTS USING IMAGE J SOFTWARE Ashleigh McLean – Bennett Lab (NRL): Revised: 2011 03 12 Online help http://rsb.info.nih.gov/ij/docs/index.html Image J for microscopy analysis http://www.macbiophotonics.ca/imagej/ Download for Mac and PC online http://rsbweb.nih.gov/ij/

1. Open program. Check to see if you are working with the latest update by going to the help menu and choosing update Image J. Follow update instructions.

2. Open the tif file of gel. The files must be in grayscale. The file must contain the experimental blot and the actin blot side by side (not one on top of the other).

NOTES: The photo should contain the entire lane and a sizable portion of the blot above and below the bands of interest or the entire blot itself if your band of interest and loading controls, for example, are on the same gel or if you wish to quantitate multiple bands on a single gel. Do not crop too tightly around your bands of interest if you choose to analyze only part of your gel (see figures below). Never adjust levels or contrast in photoshop or other graphic editing program prior to analysis with Image J. The exposure and the background of your blots are the most important issues to consider. The band intensities must be within linear range of the dynamic range of your membrane exposure. If your bands are too saturated then you cannot perform densitometry and you must repeat your experiment. Note this applies to both your experimental blot and your loading control. If your loading control is out of linear range, you must strip and reprobe your blot to obtain an optimal exposure before proceeding. Examples are provided below.

a) optimal exposure for quantification

b) A quantifiable exposure of both experimental and loading control blots

c) exposures reaching the limit of quantification given exposure and background issues (lane 3 for example in the experimental blot is saturated as is lanes 4 and 5 in the actin blots). We recognize that many researchers accept these saturation levels as the extreme limit of tolerance they use for densitometry. We will analyze this gel in this protocol example recognizing these realities of Western blotting, however we would like to emphasize that accuracy depends upon exposure and background. Blots (a) and (b) represent the level of rigour in immunoblotting that should be obtained for densitometry. This blot represents the most extreme outlier that could be included in replicate or triplicate measurement.

d) exposures in which multiple lanes are saturated and cannot be accurately quantified. Blots at these exposure levels should never be used for densitometry.

3. Under the Analyze menu, choose calibrate and open the calibration.txt file (note black=0 and white =256). This file will convert 0 to black and 256 to white. If this file is not on your computer then create it:

Column Left Right 0.0 100 46 75 105 50 175 25 252 0

4. Next, choose function “straight line” in the pull down menu of the calibration pop up window. (You may have to enlarge the window by pulling on the lower right corner to see the OK button). Click OK

5. A standard curve will appear.

6. If you would like to see the calibration bar, click on the image and then choose Analyze, Tools, Calibration bar. This will give you a new image with the gray scale values used to calibrate the gels. This image can be saved for use in a presentation describing how quantitation was performed if desired. It also shows you what is in linear range.

7. Click back on your image you would like analyzed. Click on the tool bar (usually

in top right corner of your screen labeled Image J with boxes and circles etc) and choose rectangular selection. You can choose to measure one band or multiple bands in the same lane at the same time.

To decide width: Examine your ROI and lane size. Make sure the box is wide enough to cover the largest band and will not overlap with a band in the next lane. To decide height: Create a box with maximum height possible (above and below your band of interest) that does not include other non-specific bands if possible. If a second band is included, you must examine your profile carefully to establish accurate background levels above and below the band of interest.

Note: for more accurate assessment, you can zoom into your image under the Image menu, choose Zoom > In. If you resize this box by accident, you must choose gels and reset and start again.

8. Move this box to the first lane. Under the Analyze menu, choose gel and “select first lane” (or press apple control 1).

9. Nudge to the next lane and under the Analyze menu, choose gel and “select next lane” (or press apple control 2). Repeat until all lanes have been chosen (selecting by pressing apple control 2 for each lane).

Note: if the second box automatically moves back over to the first lane, the height your box needs to be increased. Reset and increase the height of the initial box. Your height is too small to be considered an area of interest.

10. Under the Analyze menu, choose gel and “plot lanes” (or press apple control 3 on

a Mac). Lane profile plots will appear. Use the hand tool to scroll down the page and see each plot.

If your curves do not have a beginning or end, do it again. Verify that your curves appear as a hill (A) and not a valley (B). If peaks are appearing as a valley, under the Analyze menu, choose Gels > Gel Analyzer Options… remove check mark from ‘invert peaks’ option and replot.

A) B)

You cannot quantify gels that are not within linear range or where background is too high such that you cannot view a complete peak. If bands are saturated/overexposed (C – left band: saturated, right band: optimal), blots should be reprocessed. If saturated, plot will be round and not accurate for quantification (D top). Optimal plot should form a symmetrical peak (D bottom).

C) D)

11. Use the line tool to draw a straight horizontal line (hold shift key down to constrain lines to be vertical) defining the x-axis (the background). The line must touch both sides of the curve. Repeat for each lane to be analyzed.

Note: Use common sense to determine where the line is drawn – the area BELOW the line is the background. After creating the horizontal lines for all peaks, compare and evaluate the areas you have excluded in comparison to the background intensity on the tif for each lane.

12. Drawn a vertical line from the top of the curve (usually at the peak, but may be slightly skewed to the left or right) down to the horizontal line you created in step 11. Note: After each vertical line you draw, look at the whole curve and where you cut it in half - each half should be relatively as equal as possible. If not, under the Edit menu, choose Undo to erase the line and redraw it until both sides appear equal.

13. Measure the area under the curve (A) by clicking inside the shape on left or right side* defined by the line and the curve using the wand tool.

A) B)

The results window will appear with the area measurements (B). These can be saved as a text file and exported into excel or copied and pasted into excel. * How to select which side to measure When background is present above your band of interest (C, band on left) you will need to measure the area under the right side of your peak to get accurate values (D, top). It is preferable to use the same side of the curve throughout the analysis of the blot (i.e., area under right side of peak for every peak OR area under left side of peak for every peak).

If your blot is clean (C, band of right) the peak produced should be symmetrical, therefore either side should give you similar values (D, bottom). In western blotting, this is not always the occurrence – in which case use your best judgment to choose the side that best represents the band intensity in the tif.

C) D) Note: You may see “error?” beside the number – this indicates that there are too many pixels in your band that reach or exceed the maximum and/or minimum value and the program is indicating you may have reached the limit of sensitivity or that the backgrounds vary significantly between lanes– you should repeat the experiment get a better exposure. There are exceptions to this rule based on your experimental question. For example, if protein expression is reduced below the limit of detection on a western blot by the experimental manipulation (i.e., a time course following injury) in an exposure wherein all the other time points are within linear range for quantification (and thus you get the “error” message when assessing this lane as there is no quantifiable peak) then in your graph you should report this lane as below the limit of detection.

14. When reporting your use of Image J, you should state, “Densitometry was performed using Image J analysis software (NIH)” as per the request of the Image J developers.

15. If you want to do multiple blots, close the results window, reset the lane counter in Analyze Gels, and choose select none under edit, selection before you will be able to get the rectangular ROI tool back and choose the first lane.

16. NOTE: you can only do one parallel set of bands at a time. If you do two bands

one below the other, the lane values are averaged and the results are artifact. Things you must consider for densitometry Standardization to actin

1. Take the ratio of each experimental lane to its actin lane (example below: Normalized = Cx32 value / Actin value)

2. Standardize each lane to the control lane to get fold increase.

NOTE: Many researchers make the error of standardizing their results to a control lane, without standardizing the control lane to itself. This means that average value of the exp/loading control ratio in each of the blots analyzed is set to 1 and all other lanes are expressed as fold-change relative to control (which is acceptable for a small sample size as shown above) but there is no variance/error bars reported for the control lane. This can lead to false positive reporting of statistical significance. Should you wish to provide error bars for your controls, you calculate the mean ratio exp/loading control ratio of each control lane. You take the average of this ratio. You then divide each individual ratio (control AND experimental lanes) by this average. Your reported results are the mean and SEM or SD of these calculations. Your control lane will still have a mean of 1 but you will also accurately reflect the reproducibility of this control with error bars around this mean of 1. This will allow you to perform accurate statistical analysis.