Measuring Fluorescence density of nuclei

[Anoekvanleeuwen]

Hi,

I have a question about measuring the amount of fluorescence with ImageJ.
I stained my nuclei with DAPI and I'm trying to measure the fluorescence density, and with this the amount of DNA.

First, I used a method to calculate the CTCF, where I drew a circle around the nucleus, and with CTRL + M I measured the area, mean grey value and Integrated Density. Unfortunately I found out that the size of the circle had influence on the CTCF, which in my opinion is not right.

Then I heard of a 'threshold' method. I tried to find more about this method, but the only thing I can find is that I should use the option Image > Adjust > Threshold. This sounds fine, but how do I know what setting I should use? And when I know the settings, should I just measure the whole picture? And after this, is the Integraded Density my outcome or should I use a calculation?

Another question, if I'm using the threshold method, how can I do this the easiest/fastest?
I have around 1500 pictures each containing one or two nuclei.

Thanks in advance!
Anoek

[Mark Hiner-2]

Hi Anoek,

This sounds fine, but how do I know what setting I should use?
>

If you run "Image > Adjust > Auto Threshold" and use the "Try all" method,
you will get a montage that displays the results of all the threshold
methods. You can use this result on a few sample images to guide your
selection. I suggest reading the wiki page for more information on the
individual threshold methods: http://fiji.sc/Auto_Threshold

Another question, if I'm using the threshold method, how can I do this the
> easiest/fastest?
>

You could certainly write a macro that thresholds and measures all of your
images. Take a look at the tutorial for macro programming:
http://fiji.sc/Introduction_into_Macro_Programming

Basically what you'll want to do is:
1) use the "Try all" auto thresholding to choose a threshold method
2) use the macro recorder (Plugins > Macros > Record...) to record the
analysis steps for a single image
3) adapt the Process_Folder template to apply your macro to all the images
in a directory.

You can access the Process_Folder template when you have a macro open in
the editor (e.g. via Plugins > New > Macro) by using the menu entry:
Templates > IJ1 Macro > Process Folder

And when I know the settings, should I just measure the whole picture?
>

The goal of thresholding here is to create a region of measurement
corresponding to the nuclei, so you wouldn't be measuring your complete
image.

And after this, is the Integraded Density my outcome or should I use a
> calculation?
>

I presume that you just want to measure the nuclei regions established via
thresholding. You can adjust what exactly is being measured via the Analyze
> Set Measurements option. You can read more about what each measurement
option represents here:
http://rsbweb.nih.gov/ij/docs/guide/146-30.html#sub:Set-Measurements...

Hope that helps!
- Mark


On Tue, Jun 10, 2014 at 1:04 PM, Anoekvanleeuwen <
[hidden email]> wrote:

> Hi,
>
> I have a question about measuring the amount of fluorescence with ImageJ.
> I stained my nuclei with DAPI and I'm trying to measure the fluorescence
> density, and with this the amount of DNA.
>
> First, I used a method to calculate the CTCF, where I drew a circle around
> the nucleus, and with CTRL + M I measured the area, mean grey value and
> Integrated Density. Unfortunately I found out that the size of the circle
> had influence on the CTCF, which in my opinion is not right.
>
> Then I heard of a 'threshold' method. I tried to find more about this
> method, but the only thing I can find is that I should use the option Image
> > Adjust > Threshold. This sounds fine, but how do I know what setting I
> should use? And when I know the settings, should I just measure the whole
> picture? And after this, is the Integraded Density my outcome or should I
> use a calculation?
>
> Another question, if I'm using the threshold method, how can I do this the
> easiest/fastest?
> I have around 1500 pictures each containing one or two nuclei.
>
> Thanks in advance!
> Anoek
>
>
>
> --
> View this message in context:
> http://imagej.1557.x6.nabble.com/Measuring-Fluorescence-density-of-nuclei-tp5008116.html
> Sent from the ImageJ mailing list archive at Nabble.com.
>
> --
> ImageJ mailing list: http://imagej.nih.gov/ij/list.html
>

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[nmichae]

Hi,

I was also using the same function for measuring the density of activated area in the brain. What I did is, I kept the area chosen constant between the different images, using the command (ctrl+shift+E). This transferred the same ROI to a different image so that I could measure the density of the same area in different images using ctrl+M command afterwards.
But now I have an issue. What is actually the numner which we are getting when we give measure command? if the number which we get is directly proportional to the density of the ROI, it is not matching. Beacuse where I expect higher density (clear even with mere eye inspection), I get a smaller number where as where there lower density I get a high value. So what am I actually calculating? Im chosing -- set measurements--mean grey value.

regards,

Neethu

 

[Olivier Burri]

Hi Neethu and all,

First a couple of questions:
 - What kind of microscope are you using? CONFOCALS are usually a no-go for 'fluorescence intensity' measurements. If all the cells are acquired at the same distance from the coverslip and the same volume is taken, you might have a chance, but I have not yet seen data confirming this.
- Have you kept all imaging and sample preparation parameters identical over each of your 1500 images?
- How were your stainings done?

As a good starter, I'd suggest you have a quick read at:

*Seeing is believing? A beginners' guide to practical pitfalls in image acquisition*
http://jcb.rupress.org/content/172/1/9.full
The following paragraph should give you some information:
*Quantification of images—why is it useful and when is it appropriate?*

Fluorescence signal is very linear in WIDEFIELD microscopy and correlates well with fluorophore concentration, up to a point which makes it OK for semi-quantitative or relative measurements. If you expect a 5 fold or more difference then it should be OK.

Also remember that DAPI only stains AT and not GC, so depending on your biological question, this could affect any interpretation you would like to draw from your experiments. And with a quick search I have not found any papers claiming that light microscopy yields information on DNA concentration from DAPI stainings. The closest I have found, and am likely to believe, is using FACS.
*Critical Aspects in Analysis of Cellular DNA Content*
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2976661/

Also, when working on thresholds to measure intensities, it’s good practice to use a 'neutral' stain to detect the area. Setting a threshold on the intensity to then measure that same intensity biases the results. One stain marks the area independently of the stain from which you would like to extract the intensity information

In the end, fluorescence microscopy like the one you are performing is good for answering "Where is this?", not so much for "How much of this is there?"

Finally, if you still want to go about with this, an idea would be something in the lines of:

1. Blur your image significantly to smooth out uneven DAPI patches.

2. Apply a "Find maxima" filter to get a single point per nucleus.

3. Enlarge these points to get a surface of a fixed size that fits inside the smallest nucleus you can find.

Measure these surfaces (No threshold) on your non-blurred image (You can also measure it in your blurred image). Integrated density and average density should yield the same when you divide the integrated density by the area.

Make sure you have controls and/or at least two conditions to compare! Just absolute differences will mean nothing unless you have something to compare them to. Something like fibroblasts vs. tumor cells where you KNOW you will get a difference. You can thus also assess the natural variation to expect from a given sample.

This was by no means a message to discourage you, but just to inform you of the potential dangers and critiques that using DAPI quantification could bring.

All the best!

Oli

Olivier Burri
Engineer - Image Processing
& Software Development
EPFL - SV - PTECH - PTBIOP

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[nmichae]

Hello Oliver,

Thank you for the message. But what Im doing is not fluroescence measurement but we do optical imaging. and we get the neuronal activity in a grey scale activity map. So my intention was to measure the density of this activity patch. So where i expect a high density or where I have a darker activity patch, which denotes high brain activity, I 'm getting a small measurement value when I use this tool in imageJ. Thats why I was wondering, what actually is being measured here. Im very bad in explaining. Dont know whether you will understand what Im trying to say. 

Regards,

Neethu

On Thu, Jul 24, 2014 at 10:28 AM, Olivier Burri [via ImageJ] <[hidden email]> wrote:

Hi Neethu and all,

First a couple of questions:
 - What kind of microscope are you using? CONFOCALS are usually a no-go for 'fluorescence intensity' measurements. If all the cells are acquired at the same distance from the coverslip and the same volume is taken, you might have a chance, but I have not yet seen data confirming this.
- Have you kept all imaging and sample preparation parameters identical over each of your 1500 images?
- How were your stainings done?

As a good starter, I'd suggest you have a quick read at:

*Seeing is believing? A beginners' guide to practical pitfalls in image acquisition*
http://jcb.rupress.org/content/172/1/9.full
The following paragraph should give you some information:
*Quantification of images—why is it useful and when is it appropriate?*

Fluorescence signal is very linear in WIDEFIELD microscopy and correlates well with fluorophore concentration, up to a point which makes it OK for semi-quantitative or relative measurements. If you expect a 5 fold or more difference then it should be OK.

Also remember that DAPI only stains AT and not GC, so depending on your biological question, this could affect any interpretation you would like to draw from your experiments. And with a quick search I have not found any papers claiming that light microscopy yields information on DNA concentration from DAPI stainings. The closest I have found, and am likely to believe, is using FACS.
*Critical Aspects in Analysis of Cellular DNA Content*
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2976661/

Also, when working on thresholds to measure intensities, it’s good practice to use a 'neutral' stain to detect the area. Setting a threshold on the intensity to then measure that same intensity biases the results. One stain marks the area independently of the stain from which you would like to extract the intensity information

In the end, fluorescence microscopy like the one you are performing is good for answering "Where is this?", not so much for "How much of this is there?"

Finally, if you still want to go about with this, an idea would be something in the lines of:

1. Blur your image significantly to smooth out uneven DAPI patches.

2. Apply a "Find maxima" filter to get a single point per nucleus.

3. Enlarge these points to get a surface of a fixed size that fits inside the smallest nucleus you can find.

Measure these surfaces (No threshold) on your non-blurred image (You can also measure it in your blurred image). Integrated density and average density should yield the same when you divide the integrated density by the area.

Make sure you have controls and/or at least two conditions to compare! Just absolute differences will mean nothing unless you have something to compare them to. Something like fibroblasts vs. tumor cells where you KNOW you will get a difference. You can thus also assess the natural variation to expect from a given sample.

This was by no means a message to discourage you, but just to inform you of the potential dangers and critiques that using DAPI quantification could bring.

All the best!

Oli

Olivier Burri
Engineer - Image Processing
& Software Development
EPFL - SV - PTECH - PTBIOP

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[nmichae]

Hello Oliver,

Thank you for the message. But what Im doing is not fluroescence measurement but we do optical imaging. and we get the neuronal activity in a grey scale activity map. So my intention was to measure the density of this activity patch. So where i expect a high density or where I have a darker activity patch, which denotes high brain activity, I 'm getting a small measurement value when I use this tool in imageJ. Thats why I was wondering, what actually is being measured here. Im very bad in explaining. Dont know whether you will understand what Im trying to say.

Regards,

Neethu

[Olivier Burri]

Hi Neethu,

The reason we were talking about fluorescence came from the title of the email... If this is a different question, it would be advisable to change it.

I am still unsure on what you mean by optical imaging and activity maps. Is the activity map an image or some computer rendition of another acquisition tool, like EEG data? The best approach would be for you to provide an example image....

Otherwise, a remark: If the areas with higher density are dark, then it would make sense that the mean value is small, no? Dark: low values and bright:high values.

If you want the values to be inverted then just use "Edit"->"Invert", then your average intensities will be higher when there is a higher density.

Good luck.



Olivier Burri
Engineer - Image Processing
& Software Development
EPFL - SV - PTECH - PTBIOP

> -----Original Message-----
> From: ImageJ Interest Group [mailto:[hidden email]] On Behalf Of
> nmichae
> Sent: jeudi 24 juillet 2014 11:48
> To: [hidden email]
> Subject: Re: Measuring Fluorescence density of nuclei
>
> Hello Oliver,
>
> Thank you for the message. But what Im doing is not fluroescence measurement
> but we do optical imaging. and we get the neuronal activity in a grey scale
> activity map. So my intention was to measure the density of this activity patch.
> So where i expect a high density or where I have a darker activity patch, which
> denotes high brain activity, I 'm getting a small measurement value when I use
> this tool in imageJ. Thats why I was wondering, what actually is being measured
> here. Im very bad in explaining.
> Dont know whether you will understand what Im trying to say.
>
> Regards,
>
> Neethu
>
>
> On Thu, Jul 24, 2014 at 10:28 AM, Olivier Burri [via ImageJ] <
> [hidden email]> wrote:
>
> > Hi Neethu and all,
> >
> > First a couple of questions:
> >  - What kind of microscope are you using? CONFOCALS are usually a
> > no-go for 'fluorescence intensity' measurements. If all the cells are
> > acquired at the same distance from the coverslip and the same volume
> > is taken, you might have a chance, but I have not yet seen data confirming
> this.
> > - Have you kept all imaging and sample preparation parameters
> > identical over each of your 1500 images?
> > - How were your stainings done?
> >
> > As a good starter, I'd suggest you have a quick read at:
> >
> > *Seeing is believing? A beginners' guide to practical pitfalls in
> > image
> > acquisition*
> > http://jcb.rupress.org/content/172/1/9.full
> > The following paragraph should give you some information:
> > *Quantification of images—why is it useful and when is it
> > appropriate?*
> >
> > Fluorescence signal is very linear in WIDEFIELD microscopy and
> > correlates well with fluorophore concentration, up to a point which
> > makes it OK for semi-quantitative or relative measurements. If you
> > expect a 5 fold or more difference then it should be OK.
> >
> > Also remember that DAPI only stains AT and not GC, so depending on
> > your biological question, this could affect any interpretation you
> > would like to draw from your experiments. And with a quick search I
> > have not found any papers claiming that light microscopy yields
> > information on DNA concentration from DAPI stainings. The closest I
> > have found, and am likely to believe, is using FACS.
> > *Critical Aspects in Analysis of Cellular DNA Content*
> > http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2976661/
> >
> > Also, when working on thresholds to measure intensities, it’s good
> > practice to use a 'neutral' stain to detect the area. Setting a
> > threshold on the intensity to then measure that same intensity biases the
> results.
> > One stain marks the area independently of the stain from which you
> > would like to extract the intensity information
> >
> > In the end, fluorescence microscopy like the one you are performing is
> > good for answering "Where is this?", not so much for "How much of this
> > is there?"
> >
> > Finally, if you still want to go about with this, an idea would be
> > something in the lines of:
> >
> > 1. Blur your image significantly to smooth out uneven DAPI patches.
> >
> > 2. Apply a "Find maxima" filter to get a single point per nucleus.
> >
> > 3. Enlarge these points to get a surface of a fixed size that fits
> > inside the smallest nucleus you can find.
> >
> > Measure these surfaces (No threshold) on your non-blurred image (You
> > can also measure it in your blurred image). Integrated density and
> > average density should yield the same when you divide the integrated
> > density by the area.
> >
> > Make sure you have controls and/or at least two conditions to compare!
> > Just absolute differences will mean nothing unless you have something
> > to compare them to. Something like fibroblasts vs. tumor cells where
> > you KNOW you will get a difference. You can thus also assess the
> > natural variation to expect from a given sample.
> >
> > This was by no means a message to discourage you, but just to inform
> > you of the potential dangers and critiques that using DAPI
> > quantification could bring.
> >
> > All the best!
> >
> > Oli
> >
> > Olivier Burri
> > Engineer - Image Processing
> > & Software Development
> > EPFL - SV - PTECH - PTBIOP
> >
> > --
> > ImageJ mailing list: http://imagej.nih.gov/ij/list.html
> >
> >
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> > discussion
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