Shading correction for different objectives and channels

I’ve finished my testing of concentrated dye solutions for flat-fielding images. As described previously (1, 2), we’re using concentrated dye solutions to collect shading correction images, following the work of Michael Model. Following his protocol, we use 100 mg/ml fluorescein, rose bengal, and acid blue 9 for correcting the FITC, Cy3, and Cy5 channels, respectively. Additionally, we’ve found that 50 mg/ml 7-diethylamino-4-methylcoumarin is a good dye for collecting shading images for the DAPI channel.

A detailed protocol for collecting the shading images is posted on the NIC wiki, but in brief we first collect a dark image with no light going to the camera, and then collect multiple images of each dye at different positions, and calculate the median of these images to eliminate any spatial nonuniformities (e.g. dust particles) in the dye itself. Example dark and flat-field images are shown below.

Darkfield FITC_10x

We have measured flat-field images for the DAPI, FITC, Cy3, and Cy5 objectives, and for 10x / 0.45, 20x / 0.75, 40x / 0.95, and 100x / 1.4 objectives. The resulting flat-field images are compared in the montage below:


Montage of flat-field images from the indicated objectives and channels.

The correction images don’t vary as much from channel-to-channel as they do from objective-to-objective, and interestingly, the 100x objective appears to be flatter than the lower magnification objectives. We can further quantitate this by comparing the deviation from a uniform image for each flat-field image as well as the deviation from a mean image for each channel or a mean image for each objective:


Left: root-mean square (RMS) difference of each flat-field image from a uniform image. Larger values indicate higher deviation (more shading). Center: RMS diffrerence of each flat-field image from the mean image for that channel; difference is multiplied by 2. Right: RMS difference of each flat-field image from a mean image for that objective; difference is multiplied by 2.

As can be seen, the objective has a bigger effect on the shading correction than the channel. The mean error when using a single correction per channel is 5.2%; using a single correction per objective, it is 3.8%. By contrast, using a single mean image to correct results in a mean error of 5.8%. These compare to a 15% error for no correction at all. Although incorporating both objective and channel is necessary to achieve the best correction, using a single correction for all images taken with one objective will perform pretty well.

Fortunately, recording the correction images doesn’t take too long, and the appear to be stable for many days. We haven’t yet followed the performance over time to see how often flat-field images need to be remeasured but I’m hopeful that they will be stable for months and so only need to be remeasured when the microscope is adjusted or realigned.

Finally, here are some examples of our favorite kidney section (Molecular Probes slide #3) stitched at 10x with and without correction (these have been downscaled by 50% to save space):


3 x 3 stitched image taken with 10x / 0.45 objective; no shading correction.


Same image, with shading correction.


7 thoughts on “Shading correction for different objectives and channels

  1. Hi,

    This is extremely helpful. Also was wondering if you know of any good dyes for correcting the Alexa 594 channel?

  2. Probably rose bengal is bright enough in the 594 channel to use. If not, you could try acid fuschin (the other red dye recommended by Model) or rhodamine or a red-shifted rhodamine. The main requirements are high water solubility (so you can get a solution with sufficiently high OD) and low cost. Even weakly fluorescent dyes are bright enough since they’re used at such high concentrations (and in fact at these concentrations we see a lot of self-quenching).

  3. Dear Kurt Thorn,

    I have learnt and learning a lot of valuable inputs from ibiology microscopy course and this blog. I am thankful to you and your team for great effort for educating many students worldwide.

    I have a series of questions regarding quantitative imaging.

    1. what are the precautions need to be taken to use microscope(like shading correction and all..)
    2. what is the accuracy of this technique(how much trustable the results are)? because i read in cold spring protocols as

    “In general, DNA measurements are highly variable, with a standard deviation close to half the average measurement.” (

    3. can we use DNA quantities (relative fluorescence units) collected from a single layer of cell to detect DNA quantity in a cell embedded in a tissue. please assume as both were taken in same microscope, as Z slice (for both thick and single cell layer. 20 micron thick vs 100 micron deeply buried cell).?

    4. which dyes are preferable for quantitative DNA imaging?

    it would be great if you can release a video(s) regarding addressing these type of issues, because these are regularly used and misused in many labs.

    thanks 🙂

    • HI Pardha –

      Part of the reason you don’t see these issues discussed is that doing quantitative microscopy correctly is difficult and requires a lot of attention to detail. Here are point by point responses to your questions:

      1. See and – these are two papers that discuss some of the many precautions to take when performing these kind of measurements.

      2. I think you can generally do better than 50% error, although that depends a lot on your sample and microscope, and whether you want absolute or relative accuracy. Absolute accuracy is often difficult to achieve with microscopy. The protocol you link uses an old and relatively obscure instrument – I’ve never seen a photometer of the sort they describe. We often do a basic image analysis lab where you quantify DNA content by calculating integrated DAPI intensity in the nucleus and find that you can easily resolve 2C and 4C content.

      3. I would be very careful comparing nuclear intensities taken at two different focal planes – it’s likely that the deeper nuclei will be significantly less bright due to light scattering and aberrations due to imaging deeper into the tissue. I would probably want to have an internal standard to compare to.

      4. I don’t know which dyes are best for quantitative DNA imaging, but this paper may provide some answers:

      We will definitely consider doing such a video, although I’m not sure when we’ll be filming new videos in the future.

  4. Hi Kurt,
    Thanks for the very useful information.

    Out of curiosity, are the images in the channel/objective montage displayed on the same scale? If not, you could say that the 100x objective is flatter across a greater percentage of its field of view, but it might be that the 10x objective is flatter across a larger area. Can you comment on this?


    • These are drawn without any scaling – i.e. one pixel on the camera is one pixel in the final image. If they were scaled to constant sample distance, the 10x image would be 10x larger on each axis than the 100x image, so it would be flatter over a much larger sample area.

  5. Hi,

    Thank you for your guidelines for nor uniformity image correction. It’s very interesting and useful. I have one question:

    I do take redox imaging to measure NADH and FAD using fiber optic probe measurements. While for measuring FAD fluorescence, i could able to see the shading as you have shown in gray scale image above. Even, when i take measurement with control (only PBS, no fluorophore), i could able to see the shaded images as you have shown above (second gray scale image). Now,
    1. I would like to correct the intensity of that obtained images, could you please suggest any method?.
    2. Is there anyway to get rid of that shaded intensity ( by changing optics, lens, etc.,)

    I would like to discuss more about my image outcome. my mail id is I would be happy if you could able to resolve this problem.

    Thanks once again

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