Hi there!

I have done similar studies with the mitotracker and they method I follow is listed quite nicely in the following help page from FIJI

http://fiji.sc/Colocalization_Analysis

Pearson's correlation coefficient, as you will notice is not absolute. You can also try out Manders coefficients!

Good luck with the experiments!

You can find a nice review here:

Kenneth W. Dunn, Malgorzata M. Kamocka, and John H. McDonald; A practical guide to evaluating colocalization in biological microscopy, Am J Physiol Cell Physiol 300: C723–C742, 2011.

and additional information here:

Jeremy Adler, Ingela Parmryd; Quantifying Colocalization: Thresholding, Void Voxels and the Hcoef, Plos One,  2014, Volume 9, Issue 11, e111983

Green and red make yellow. If you can clearly see difference in yellow signal between your treatment and control, the confocal images (without quantitation) are generally acceptable for publication, even in top journals.

you can use Imagej from NIH , it is a free download and an open source program, you can find there a lot of answer and plug-in for your question

Hi,  

I agree with Rongqing on the difference in visual representation within your images. In my opinion, the easiest way for you to quantify your data is to count the number of mitochondria you see in your images (control and treatment). Count the number of cells you see colocalization (yellow) as well. [I assume that the protein of interest you are looking for is only present in the mitochondria right? it cannot be found in other cellular compartments.] Determine the tissue area (or the area of your confocal image) in your samples. Then calculate the proportion or percent of mitochondria per area in each sample. Out of this proportion or percent determine the number of mitochondria colocalizing your protein of interest. when you obtain this percentage data, use Fisher's exact test to determine if there is a significant difference between them.

you can also just run a simple Student's T-test ( or anova if you have multiple treatment or time point) on the number of colocalized cells  per area (control and treatment).

Hope this is helpful.

Best of luck.  

Hello

I agree with all researchers , these a good answers

you can check out of the attachments , i hope help you

Good luck

http://www.microscopyu.com/references/colocalization.html

Hello,

I agree with Marzia di chio. ImageJ is a good open source programme (using Java technology). You can find it here: http://imagej.nih.gov/ij/

There are also lots of resources (tutorials, examples, macros etc.).

Good luck!

Most acquisition software packages also provide tools for quantifying co- localisation, allowing data to be exported into Excel. Some of those are available as light versions. You might also try out the profile function -essentially drawing an arrow across a single cell or an area of interest inside the cell. This generates a histogram which can be saved and used in presentations or publications. 

Also consider FRET as an alternative to "conventional" IF if possible. 

I agree that most software programs for confocal scopes allow to analyze colocalisation. Typically it gives yellow color on merge picture. It is a bit tricky for regular fluorescent scopes (not confocal) since software is not able to compare depth of imaging section and it can appear like co-localized proteins/structures even if they are not.

Good luck))) 

Hi,

I  suggest the Colocalization Colormap plugin

This is the reference: Jaskolski F, Mulle C, Manzoni OJ. An automated method to quantify and visualize colocalized fluorescent signals. J Neurosci Methods. 2005;

146(1):42–49

And this is a recent article by my collegue in which he used this plug- in

Int J Nanomedicine. 2015 Mar 10;10:1885-98. doi: 10.2147/IJN.S77734. eCollection 2015.Peptide gH625 enters into neuron and astrocyte cell lines and crosses the blood-brain barrier in rats.

Good luck