Measuring Cell Fluorescence using ImageJ
Image J can be downloaded for free from here .
This guide can also be downloaded as a complete PDF here:Â Measuring Cell Fluorescence using ImageJ
Here is a very simple guide for determining the level of  fluorescence in a given region (e.g nucleus)
- Select the cell of interest using any of the drawing/selection tools (i.e. rectangle, circle, polygon or freeform)
- From the Analyze menu select “set measurements”. Make sure you have AREA, INTEGRATED DENSITY and MEAN GRAY VALUE selected (the rest can be ignored).
- Now select “Measure” from the analyze menu or hit cmd+m (apple). You should now see a popup box with a stack of values for that first cell.
- Now go and select a region next to your cell that has no fluroence, this will be your background.
NB: the size is not important. If you want to be super accurate here take 3+ selections from around the cell. - Repeat this step for the other cells in the field of view that you want to measure.
- Once you have finished, select all the data in the Results window, and copy (cmd+c) and paste (cmd+v) into a new excel worksheet (or similar program)
- Use this formula to calculate the corrected total cell fluorescence (CTCF).
NB: You can use excel to perform this calculation for you.CTCF = Integrated Density – (Area of selected cell  X Mean fluorescence of background readings)
- Make a graph and your done. Notice that in this example that the rounded up mitotic cell appears to have a much higher level of staining, but this is actually due to its smaller size, which concentrates the staining in a smaller space. So if you just used the raw integrated density you would have data suggesting that the flattened cell has less staining then the rounded up one, when in reality they have a similar level of fluorescence.
How to Cite this if you wold like to:
I used this method in this paper here:
Burgess A, Vigneron S, Brioudes E, Labbé J-C, Lorca T & Castro A (2010) Loss of human Greatwall results in G2 arrest and multiple mitotic defects due to deregulation of the cyclin B-Cdc2/PP2A balance. Proc Natl Acad Sci USA 107: 12564–12569
But you can also find a similar method published here:
Gavet O & Pines J (2010) Progressive activation of CyclinB1-Cdk1 coordinates entry to mitosis. Dev Cell 18: 533-543
And here:
Potapova TA, Sivakumar S, Flynn JN, Li R & Gorbsky GJ (2011) Mitotic progression becomes irreversible in prometaphase and collapses when Wee1 and Cdc25 are inhibited. Mol Biol Cell 22: 1191–1206
And my apologies to any others that I have not mentioned.
Hi, I find your post very relevant for my project. However, my cell nuclei were stained by DAPI, and my protein-of-interest was stained with Alexa Fluor 488. I am trying to quantify the amount of this protein within the nucleus. Could you give me some advice on this? Thanks!
Hi, this method works with any flurophore, be that DAPI, 488, 555, FITC, Texas Red etc etc. If you want to only measure the nucleus of a cell, then when you draw your region, just limit it to the nucleus. This method is good for gathering data on individual cells, and I normally find that a minimum of 25 is needed to get any meaningful stats (but more cells is always better). If your more interested in the whole population, then you might be better off using a threshold method which could give you the average intensity across the whole field very quickly. Just make sure what ever you do you use identical acquisition settings for every sample. Overall, quantifying IF images is perhaps a little subjective and open to user input, but it can still be a useful tool. I hope that this is of some help. If you still need further clarification let me know.
But in my case, should I separate the RGB stack into individual fields, and then measure the green and blue channel staining intensity separately? With blue channel of DAPI alone staining as background, and then use the formula as shown by you?
Thanks for the help, I am a complete beginner in this!
Hi, you need to measure each channel separately. The background measurement should be taken from an empty region on the same channel, just adjacent to the sample of interest, in your case the nucleus. You can, then also measure the DAPI staining (in the same way) if you like as well, but I’m not sure if this is really necessary. If your only interested in determining if your protein (488 channel) increases/decreases…then you only need to measure that channel. So in summary yes you need to separate the channels, and measure them independently! Good luck
Hi, in case you are interested in the whole cell population, how exactly would you do the threshold method? Thanks!
Is there a way to use ImageJ to calculate the mean fluorescence in a given field? Or will it only work for single cells?
You could use this same method but instead of selecting a single cell, you could select the whole field using for e.g. the rectangle tool.
how doyou change your RGB picture to 16bit pixels only picture?
This is so useful! Thank you very much!
Hi I am a complete beginner at using imageJ. Would I be able to use this method for comparing the intensity of alexa flurophore 488 inside the nucleus with the intensity at the periphery of the cell in order to see if the protein had moved to a different location in the cell? If so how would I go about doing this?
Any help would be appreciated. Thanks
Hi Dani, yes you should be able to use this method to do what you want… just include an extra step for the cell periphery. my suggestion is to try drawing around the total cell and then the nuclear fraction, take reading for both, correct each for background as per formulae and then divide one by the other to create a nuclear:cytoplasmic ratio, which over say 20+ samples you hopefully will be able to get some statistical relevance.
Good luck
Andrew
Hi Dani,
Have you done your measurment? I am a beginner with imageJ and have almost the same situation, could you please help me by telling the way you have used to do your measurments.
Thanks a lot,
Saeid
Hi Andrew, I wanted to ask if you have ever used size exclusion when doing your analysis in ImageJ. If you have, how do you set the scale for your maximum size?
Hi Kelly, sorry I have not done any size exclusion with image J so I’m afraid I can’t offer any advice. Good luck
Andrew
Hi Andrew,
I was using Imaris before to quantify my confocal images and beside the value of the fluorescence of a defined area, it was giving me also the size of the area, is it possible to evaluate that in Image J? (by giving him a scale in um before of course).
Thanks in advance
Cedric
Yes you should be able to do this in imageJ. First set the scale of the image by going to the menu item Image>Properties.
Then go set the type of things you want measured by going to the menu Analyze>Set Measurements. Here you can select a whole heap of different parameters to measure area being one of them !
Good luck
Andrew
this is what I searching for, thanks for sharing!
Greeting from Taiwan
Cathy
Hi Andrew. Thank you so much for your helpful post. I am a beginner with ImageJ, so I was hoping you could answer a question for me. My cells are stained with Bodipy (specific to lipids) and DAPI for the nucleus. I’m trying to measure the average fluorescence of the lipid droplets in the cytoplasm. How do I subtract the fluorescence emitted by the DAPI? Thanks!
You should do the quantification on the image before you compile all the channels together. Most microscopes should acquire each channel as a separate image. If you have a good microscope then it should cut out most of the bleed thru, also I find acquiring the Dapi last helpful, as the blue light can often bleach the other channels a bit.
Hope this helps
Andrew
Thanks! That does help a lot! I have the green fluorescence images, so I should be able to do that. Do I need to convert the image to an 8-bit black and white prior to starting the measurements? Also, if all the cells are different sizes, do I have to subtract the area of the nucleus since I’m only interested in what’s in the cytoplasm?
Not sure it has been along time since I came across a camera that took colour pics, most are greyscale. It should be fairly simple to test if conversion makes a difference to the reading but my guess is that it won’t. As for the nucleus subtraction that is really up to what it is exactly that you want to measure. I would start with the simplest setup and see if the data corresponds to what you see visually. Then add complexity if needed to improve the readout quality if needed.
Dear Marli,
I had the situation here, i also want to count the lipid droplets, did you get through with that? Please give me your suggestion, thank you
Lam Nguyen
Hi Andrew,
I found your article very useful, thanks a lot for posting on this topic! I’m also a beginner in ImageJ.. I would like to calculate nulcear/cytoplasmic fluorescence intensity ratios on my cells. I suppose, I’ll use a simple circle tool, and put down circles in the cytoplasm and in the nucleus in case of many cells and than calculate the average. This might be very simple, but I don’t see how I could set the size of the circle tool to be constant. Otherwise I need to create circles by hand all the time, and if the size is not the same, then the whole thing will get far away from being representative..
Thank you so much for the answer!
I have not yet figured out how to set a constant size, but there are two work arounds. First you can move the circle that you set up on the first image around the first image and take all your readings. Next copy and paste that circle onto the next image, then hit the delete (backspace) and it should just delete the pasted image but leave the circle, so you can then repeat the same process.
This is ok for a few images but its easy to accidentally delete or loss the circle tool.
Thus I think you maybe better off, making the readings independent of size by using the formulae in the guide to correct for both background and size of the area that you select. This way can draw any size circle, or even perhaps select the whole cell, and then then subtract the nucleus. I suggest having a bit of play to see which method/s give you the best/most accurate looking data.
Good luck
Andrew
You can achieve a constant size of your circle by running the ImageJ Macro “Circle Tool”. Then you can specify a set/constant circle size (20 pixels or whatever) and every time you click anywhere in your picture, your circle size will always be the same.
Hey, great article. Just had a quick question, I am comparing fluorescence between different samples and using the entire field. So the area fr each sample is the same. When I go to calculate my corrected total cell fluorescence, i get some negative values. Is there a way I can adjust for this?
Hi, this method is specifically designed for single cell analysis, so applying it to a whole field probably won’t work and thus the negative values. If you just want a whole field then you are better off using a threshold over the whole field as this will be faster and probably more accurate… but won’t give you individual cell info.
Good luck
a
I am a beginner in imageJ and i have a question. Am I able to use this to quantify the fluorescence in a given field? I am not interested in individual cell data. Will this method be accurate and if not do you have any suggestions on what I could do?
Hi,
This method is specifically targeted at single cell analysis, and is probably not the best method for whole field quantification. I would look into the use of thresholds to do what you want to do.
Unfortunately I am not able to help you out with that method as I don’t know how to do it…sorry but good luck
a
Hi Andrew, thanks for your post. It’s very useful. In my case I want to compare the intensity of my interested protein which is localized at the septum between wild type and mutant strain. The area of selected is very small approximate 12-15. Therefore, should I keep the same size of area in each cells ? There is much difference between multiply by 12 or 15. Or do you have any suggestion?
Thanks in advance
Kanya
hi, I am a new user of image j. I want to analyse the fluorescence intensity of two cells. but while following your procedure, I can not find the “set measurement ” option under analyse menu. please help
Yes as long as all the images are acquired using the exact same conditions then you can use this to quantify 2 or more different pictures.
hi, i have another question. can this procedure be applied for calculating intensity from two separate images
I want to measure fluorescent intensity in tissue. Do I measure small areas of the tissue and then calculate the CTCF for each one of them? The values I am getting are double what I would expect.
Thanks
G
Hi,
I was wondering if you change the LUT levels of the images, does the integrated density or the mean fluorescence intensity changes? Or are the LUTs just to enhance the contrast in the image
I normally do not change or alter the images at all. This is the best way to ensure that you don’t introduce any artefacts into your measurements. Its also critical that you acquire all your images with identical exposure times etc.
However, i’m pretty sure that altering the LUT levels will not have an effect on measurement readings. You can probably try this for yourself, take a reading pre and post and see if it changes things.
a
Thank you for the reply. I was also wondering if I want to compare multiple images for fluorescence intensities, should the area of the cells be the same?
Thanks for the informative article. I’m trying to use imageJ to quantify intensity of primary cilia in two different channels. In this case, i would like to keep the area I have selected constant between the two channels while measuring intensity of each color separately. Do you have any idea on how to do this? Thanks!
Hi you can copy and paste an area between two images. You just have to hit the back space key ‘once’ when you paste into the new image. I have not figured out a better way and if you have a lot of images then it can be very annoying when you make a mistake. Thats why I developed this method to account for differences in area size between to images.
Hi,
I, like everyone else, am new to Image J. My situation is a bit different than others. I have cells that i am co-localizing microtubules and virus. I have individual images from the two different channels, however I want to get the intensity of the areas where they are merged so I need to have both channels together. Is there any way I can only quantify the yellow areas.
Hi Lucy,
Unfortunately this method is not suitable for what you want to do. You need to look at co-localization. There are plenty of plugins for ImageJ that should help you do this. A quick google pulls up several. Also many other microscopy software programs now have co-localization built in.
good luck
a
HI, Can this method be used to compare the fluorescence intensity to find the ratio between YFP and CFP and to calculate FRET with single cell images taken through blue channel and yellow channel separately.
It can take a look at the following papers for the method.
Gavet, O., & Pines, J. (2010). Activation of cyclin B1-Cdk1 synchronizes events in the nucleus and the cytoplasm at mitosis. The Journal of cell biology, 189(2), 247–259. doi:10.1083/jcb.200909144
Gavet, O., & Pines, J. (2010). Progressive activation of CyclinB1-Cdk1 coordinates entry to mitosis. Developmental cell, 18(4), 533–543. doi:10.1016/j.devcel.2010.02.013
Although there are several FRET specific plugins [Link] for ImageJ which maybe worth a look at as this could simply the process.
a
Hi, I found your method very interesting. I’m trying to measure different fluorescence from tyrosine phosphorylated cells. My only concern is that when I apply your formula for adjusted intensity, it comes out a negative number… my areas are usually around 4000 and the back intensity around 5000.. Any idea?
That is very strange. Not quite sure what is going on there, without seeing the images, I would guess that perhaps your images are either too over or under-exposed resulting in a high background or under-estimation of your reading of interest respectively.
a
Hi,
This was so helpful. I have quantified the intensity of my cells. But, I was wondering what will be the unit, as the digits I got as a result are so high. Thank you.
The units are arbitrary, and don’t have a physical meaning. Its all just relative
Thus I label the graphs as “Total Cell Fluorescence (Arbitrary Units)”
a
Thanks for a quick reply. I also want to measure intensity of different concentration of proteins. What should I use as a background, also should I measure background each time? the proteins I am measuring are micro arrayed as a spot on the slide.
Cheers,
The background reading should be an area close to your sample of interest. It should be taken and matched for each image and fluorescence filter. For micro arrayed dots, this method may not be the most efficient. There maybe more automated solutions out there ?
a
Hi,
your post is really helpful. for measurement in the box for image J I have IntDen and also another parameter RaWIntDen, what is that?
http://rsbweb.nih.gov/ij/docs/menus/analyze.html
Hi, your post is really helpful. Here I have some questions regarding to cell area meaurment by using ImageJ. My cells are stained by phalloidin and DAPI, and I want to meausre the area of these cells and then comparing the morphology differences between my two samples. I am currently using ‘threshold method’, however, I have find that finding the region of interesting (ROI) through adjusting the ‘threshold value’ is a littile subjective. I am wondering what is the most proper way to measure the cell area in ImageJ? and Do I need to keep the thoreshold value constant during the meaurement in order to make the results comparable between my two samples? Thanks in advance Andy
CTCF = Integrated Density – (Area of selected cell X Mean fluorescence of background readings)
In the formula above, what does ” Mean fluorescence of background readings ” mean? Is it the mean value of the background image or the integral density of background image. Little more clarity on this would be really helpful to me.
Thanks
Rahul
Normally you should take several background readings for each image, so the mean of the background readings is the mean of those.
so, i should take mean of integral densities for background images ?
Thanks
Rahul
do we take the average of the integrated density of background reading or the average of the mean gray values of background readings
Thanks
Rahul
You use the mean for background not IntDen, as you will multiply the average mean by the area of the cell to give the Background- IntDen of the cell. If you used the original IntDen reading this would be for the small circle area you selected, which is not what your wanting. Hope that helps
a
Hi, this was really helpful. I just quantified some data using this. I was wondering what the difference between integrated density and CTCF is? I have been unable to find definitions for each. I used a premade excel spread sheet that calculated this number for me.
http://rsbweb.nih.gov/ij/docs/menus/analyze.html
CTCF = Integrated density corrected against the background.
Hope this helps
a
Hi, thanks for your very helpful post. I have one doubt though. I am measuring fluorescence intensity in nuclei to compare ratios of two different proxies. I have z-stacks of the image. In order to measure the CTCF I do a Sum Slices projection. Woud it be better to do Average Intensity or Maximum Intensity projections instead.
hello, imageJ is certainly a great solution to many imaging processing techniques. I have a question, I am using a dye which accumulates in healthly mitochondria and on introducing drug the dye releases out of the mitochondria. can I use fluorescence quantification for control image against treated image? if so , how? I would be really thankful for this help.
Hi Andrew,
Thanks so much for this helpful post.
Please, I would like to ask if I can use the fluorescence intDen measurement to quantify the amount of a drug molecule (ofloxacin) in tissues.
Want to determine the absorption route of this drug molecule using sliced animal tissue.
Thanks for your help.
You could but like I said to the above post, if your not interested in single cells info, a threshold approach will be faster and easier.
Good luck
a
Hi Andrew,
Your blog is really helpful. I too need some help with imageJ to quantify fluorescence of my dye on tissue samples. I have grey scale fluorescence mosaic of my stained tissue from confocal microscope. I need to quantify the amount of dye present in the samples. Could you provide some help?
Thank you
Ash
Hi, sorry for the delay, been super busy. From the sounds of it, you maybe better off using a threshold approach. The method I posted here is really only for individual cells from numerous independent fields. If your not concerned with individual cell quantification, and just want a total field quantification then thresholding will be much faster.
Hi,
I succed to use your clear method to quantify GFP of a 2D image. Thanks for This. But I would like to quantify it in a 3D image of a cell which is perfectly spheric. I have roughly 50 to 120 stacks for a cell. Should I do the quantification stack by stack or is there any other method to do this in a volume on the 3D project of my cell ?
And I would like to know if with a 3D image of a confocal laser microscope, imageJ integrate density concidering that the image is made of pixels or does it concider that the image is made of voxels ? because in my case, a volume cannot be made of pixels, and so I cannot sum the quantification of each stack. Thanks a lot for your help.
Great question, and I wish I had a good answer. What I have done in the past and I have seen other publish this also, is to just take the best single plane/slice and quantify that. You could possibly flatten the stack (max projection) but I am not sure what algorithms are used by ImageJ to do this, and if it would be consistent enough to allow quantification. You could also try this http://imagej.nih.gov/ij/plugins/voxel-counter.html, or switch to a better (but costly) 3D program like Imaris. Sorry I cannot be of more help
a
Thanks a lot for your answer. It is approcimately what I thank. I’ve found a lab where people use imaris. I think I will try it. It is probably possible with imagJ or fiji, but as you said, the result given by this software is not sure, and for a publication, it would turn into a problem. bye. François.
Sorry, in my precedent message I wanted to say slice and not stack. thanks
Hello, I’m working with FISH images, in green fluorescence of bacteriais like E.coli (populations). This method is very nice, but i think that the threshold is more easier. I don’t know how to do it! Could you give me some advice?
Thanks
Hello ! Thanks a lot for the great guide you made !
But I’ve a little problem with imageJ. In my case, I have images with 2 differents colors and I want to determine the fluorescence intensity of a specific area for both color. The problem is that I’m not able to keep the same area between my 2 stacks…so how could I copy the area on my green stack for example and paste it to my red stack (so that the side and the position of the area are the same) ?
Sorry for my bad level in english, hope it was “understanble” enough.
Thanks !
Have you ever thought about including a little bit more than just your articles?
I mean, what you say is valuable and everything. But imagine if you added some great
graphics or video clips to give your posts more, “pop”!
Your content is excellent but with pics and videos, this site could
certainly be one of the best in its field. Fantastic blog!
Hi,
Very nice and informative blog.How to measure co-localization in cells using Image J
How do I calculate nuclear-cytoplasmic ratio in ImageJ?
hi:
I read your blog, it is very helpful for my study. in my case, I wish to get density from whole section. accroding to your instruction, I am confused about your formula:
CTCF = Integrated Density – (Area of selected cell X Mean fluorescence of background readings)
in my case, I get the mean intergrated density 2666304.833. in addition, I get background density 57145.16667. area of selected tisuse is 30406. it means that CTCF=2666304.833-30406X57145.16667?
or 2666304.833-57145.16667?
Feng
I re-read your message, I get it. thank you!
Hi, thanks for doing this for us beginners. I have a problem when using ImageJ. It is that I want to meature the fluorescense in more than thousands of pictures. So is there any methods for me to do it easier, instead of doing it one by one?
Thank you so much!
hi Andrew,
in the results window, is there a way to rename each entry? this helps me keep things organized.
Hi,
I am investigating the proliferation of my cells and am using Ki-67 marker and DAPI. By looking at my cells, Ki-67 is presented in forms of ‘dots’ in the nucleus. My question is can i still use your approach? or (to be more accurate) I need to count the fraction of cells stained with the marker? (how to do that?) Another question is about the negative control cells. I understand you have used the background analysis (to calculate the CTCF) but do I skip this when I have the negative control sample? I think, I could circle the cell/s of interest (measure their area) and subtract it from the cell/s in negative control sample to get a better result. Would you say this is a correct procedure?
BTW, very informative blog
Thank you in advance for help
Sorry for the delay, my guess is that you would be better off using a dot counting approach rather than the one I posted here. I have posted a video on how to do this using Imaris.
http://vimeo.com/mitoticlab
Hi, I am new to imagej and appreciate any help. The circle tool no longer seems to be included in the download and I can’t find it anywhere on the website of imagej. Can someone please email the macro to me or tell me where I can download it?
Hi there,
Would you please advise how to quantify fluorescence in nuclei and cytoplasm separately?
Cheers,
Bill
Draw a circle around the nuclei. Then one around the whole cell. Minus nucleus from whole cell reading for cyto fraction.
a
Hi Andrew,
Thanks for sharing this blog. I was wondering that in this formula “.CTCF = Integrated Density – (Area of selected cell X Mean fluorescence of background readings), why couldn’t we multiply with average Integrated density of the 4 background readings instead. I am trying to understand the logic behind this formula. Please share ur thoughts. Thanks,
Devi
Hi Devi, the background reading is to be applied over the size of your region of interest (i.e. cell) thus the integrated density of the background is not what you want to apply to this area. Think of it this way the IntDen = Area X Mean for your cell. You then want to subtract the background for the exact same area of your cell, therefore you just substitute in the mean(background) into the IntDen formula so that you end up essentially with 2 IntDen values one for the cell and one for the background. The CTCF is really just the IntDen(cell)- IntDen(background).
Hope that helps
a
Hi Andrew,
I applied a dye to an epidermal strip and I wanted to observe the fluorescence. I when I oppened the file with the image (the one that i got from the confocal microscope and lsm file) I saw I got three images (all of them are 8-bit) one in one i coud see the the green staining (it has written in the upper part of the image 1/3 (CH2)), the other image I could see some red spots (it was written in the upper part 2/3 (CH3)) and the last image was grey (3/3(ChD)). My question is which of these images should i use to measure the fluorescence or shoud I do something else?.
Priscilla