Thank you very much for the video. Sorry, I was reading the manual and in one part it says: "...for our deconvolved 3D stacks of beta-cell mitochondria, a block size of 1.15 to 1.25 microns and a C-value of 10 or 11 is often found to be ideal" I treated these values in deconvolved photos of mitochondria from HCT-116 cells, I have the impression that this setting leads to the elimination of part of the network, increasing the perception of network fragmentation. This seems to be resolved if I increase the block size (Ex: 2.50 microns). My question is, is there a valid method to define an appropriate "block size" in Threshold Optimize? Or it's trial and error.
@@pacmorgado3443 it varies depending on the image. Your acquisition parameters may have been different from what the manual suggests. It usually is trial and error to get the “perceived” best block size. Thanks for supporting my channel.
Great video! Super helpful! I am confused about one thing - The Per Cell analysis gives me a different total area and perimeter than the area/perimeter I get after selecting ROIs and measuring. Which area is the correct area occupied by the mitochondria?
Hi @maheenwahid3903. The total area is the sum of areas of ALL mitochondria in the image. The total perimeter is the sum of perimeters of all mitochondria in the image. The mean perimeter is the toal perimeter divided by the total number of mitochondria (count) in the image. If you do analysis on a per-Mito basis, the Area or Perimeter is the measurement for each mitochondrion.
@@johanna.m.dela-cruz okay that makes sense! And when I select all ROIs and measure area, shouldn’t that give me the sum of areas of all mitochondria too?
@@maheenwahid3903 When you have the ROIs, click Measure and you should get measurements for each mitochondrion. If you get the sum of all the areas, the value should be close to what the per-Cell measurement would give you.
Sorry, I have questions about how to adjust the distance between each image in the Z-Stack. According to an AI, I could change in Image > Properties to change the size of the "Voxel depth" and set it with my z interval, which is 0.3 microns. Is that correct? Beforehand thank you very much.
Hi! Thanks for a great tutorial. When I threshold the image using default settings, I see in the ROI manager that several (individual) mitochondria are getting counted as a single ROI leading to massive numbers in the analysis. Is there any advice on how to reduce this? I used the nyquist equation to set my scanning parameters and I do not see a difference.
@@nickdenniston4271 hello. Did you try the Optimize Threshold command first to determine the right settings for your image? Default settings work most of the time, but not everytime. You could also try deconvolving your images first before using the plugin.
@ ive deconvolved the images and i have also tried a variety of threshold setting indicated by the optimize threshold command. The best setting i found is a block size of 1um and a C value of 16. With these settings i am not getting as much noise and it minimizes the issue of having multiple mito being grouped into a single ROI. I am now worried that this C value is too high and i am missing information. Any thoughts?
Thanks for this amazing tutorial. I am trying to use this to do a ratiometric analysis using two channels. Is there a way to do it on a z-stack image and get the mean intensities in each of the channels?
Hi! Thanks for watching. If you want to do ratiometric measurements, maybe you can take a look at my other tutorial: th-cam.com/video/66NkiuTJtx4/w-d-xo.htmlsi=TTw2jk5A9ScTym1X
Halo, if i want the measure the length of mitochondria which data should I use? total branch mitochondria percell or I choose the data from the total branch length per mito? thank you
Hi, I tried several times to work with Mitochondrial analyzer function, Although I plugged in the mitochondrial analyzer but during using the 2D threshold function I am getting an error message like"marco or script not found".
Hi !@@johanna.m.dela-cruz i am also facing the same problem. I am not able to get the jar files of these three : 3D ImageJ Suite, ImageScience and IJPB-Plugins.
@@promilalakra9886 hello. To enable these sites in Fiji, go to Help › Update… Click Manage update sites, Check ImageScience, etc., Click Close. Click Apply changes Restart Fiji.
Firs of all, great tutorial (most music and text ;o). If any one can help me, I had same trouble with the delimiter/units of the 3D Sphericity output. For a sphere, it is spected values from 0 to 1, but for same mitochondria I found values beetween 0.4 to 1.065. When I converted the data for comma, which I use it for a statistic program (Prisma Graph), I dont know if Im working with 1,065 or 1065,0; which for both cases is strange since the maximum expected value is 1. So, what Mitochondria Analyzer output parameters can I use, to recalculate the sphericity and to confirm these numbers?
Hi Guilherme. Thanks for your interest. I have noticed this as well. I think that sphericity is calculated based on MorpholibJ's formula: "the ratio of V^2 by S^3, multiplied by 36*pi."
@@johanna.m.dela-cruz Thank you for the quick answear and for the equation. Now I will try to figerout how to "interpret" the number delimiter (point to comma)
if (canSigma) run("Sigma Filter Plus", "radius=" + sRadius + " use=2.0 minimum=0.2 outlier stack"..................why for thresholding in 3D, it does not recognize this command.
Hey I'm trying to measure the mitochondrial length in the same way. Although I can clearly see the difference in length between my control and test sample, still upon measurement the difference is not reflected. How do I manage that
Hello Arpita. Are you doing analysis on a per-cell or per-mito basis? If you want results for each mitochondrion in each image, you will need per-mito calculation to identify mitochondrial sub-populations. Per-cell basis should give normalized values rather than raw totals.
@@johanna.m.dela-cruz I did both. In the control vs test I can see the changes, but when measuring the mean length or length/mito i didn't see any changes My another concern is also that if I change thresholding parameters in control or in test ami I not making the data biassed?
@@arpitadutta6737 The thresholding strategy of the plugin involves a local or adaptive thresholding algorithm that requires empirical determination of its settings. This means that finding optimal settings for thresholding values is largely based on your own observations. You will have to determine which thresholding algorithm to use based on your own judgement....Quantification will change based on the block size and C-values you use. S0, whatever method and parameters you choose will have to be applied to all your images (assuming they were acquired using the same settings).
@@johanna.m.dela-cruz heya. I tried to put adaptive threshold, changing the c value to 1 also didn't show any change, also I changed the block size into 0.25 from 1.125; that is the default. Still the thresholded image does not reciprocate to the original.
@@arpitadutta6737 Thresholding is always a challenge especially with complex images. I don’t know how your actual images look like, but you may want to try pre-processing them before using the plugin. Are your images 2D or are they z stacks? Some complex images might need deep learning algorithms for more accurate segmentation.
My ImageJ crashes when I try to do the thresholding. I can't proceed further 🙁 I have tried uninstalling and re-installing ImageJ. Does anyone know what's the issue?
@johanna.m.dela-cruz the 3D image is 108mb, and the 2D image is 4 mb. I tried both 3D and 2D threshold but they both crash the app without any prompt. I am on Windows.
Hey. I had replied to the comment but somehow it didn't post and I didn't realise. Thanks for the quick response. I figured out that it was a memory issue. I allocated more memory and now it works fine. 😊
![FIJI (ImageJ): Fiber Orientation and Direction [Directionality & OrientationJ]](http://i.ytimg.com/vi/tN0bEVsumJc/mqdefault.jpg)
![FIJI (ImageJ): Fiber Orientation and Direction [Directionality & OrientationJ]](/img/tr.png)
![FIJI (ImageJ): Counting Foci or Puncta [Difference of Gaussians]](/img/n.gif)
This is fantastic. great tutorial! Thank you
Thanks, that means a lot.
Thank you very much for the video.
Sorry, I was reading the manual and in one part it says:
"...for our deconvolved 3D stacks of beta-cell mitochondria, a block size of 1.15 to 1.25 microns and a C-value of 10 or 11 is often found to be ideal"
I treated these values in deconvolved photos of mitochondria from HCT-116 cells, I have the impression that this setting leads to the elimination of part of the network, increasing the perception of network fragmentation. This seems to be resolved if I increase the block size (Ex: 2.50 microns).
My question is, is there a valid method to define an appropriate "block size" in Threshold Optimize? Or it's trial and error.
@@pacmorgado3443 it varies depending on the image. Your acquisition parameters may have been different from what the manual suggests. It usually is trial and error to get the “perceived” best block size.
Thanks for supporting my channel.
Great video! Super helpful! I am confused about one thing - The Per Cell analysis gives me a different total area and perimeter than the area/perimeter I get after selecting ROIs and measuring. Which area is the correct area occupied by the mitochondria?
Hi @maheenwahid3903. The total area is the sum of areas of ALL mitochondria in the image. The total perimeter is the sum of perimeters of all mitochondria in the image. The mean perimeter is the toal perimeter divided by the total number of mitochondria (count) in the image. If you do analysis on a per-Mito basis, the Area or Perimeter is the measurement for each mitochondrion.
@@johanna.m.dela-cruz okay that makes sense! And when I select all ROIs and measure area, shouldn’t that give me the sum of areas of all mitochondria too?
@@maheenwahid3903 it should.
@@johanna.m.dela-cruz I’m getting different areas for ROIs and Per Cell :(
Which one is cell mitochondrial area I should consider?
@@maheenwahid3903 When you have the ROIs, click Measure and you should get measurements for each mitochondrion. If you get the sum of all the areas, the value should be close to what the per-Cell measurement would give you.
Sorry, I have questions about how to adjust the distance between each image in the Z-Stack. According to an AI, I could change in Image > Properties to change the size of the "Voxel depth" and set it with my z interval, which is 0.3 microns. Is that correct? Beforehand thank you very much.
The voxel depth is basically the z interval, so if you go to Image > Properties, you can set the voxel depth from there.
Hi! Thanks for a great tutorial. When I threshold the image using default settings, I see in the ROI manager that several (individual) mitochondria are getting counted as a single ROI leading to massive numbers in the analysis. Is there any advice on how to reduce this? I used the nyquist equation to set my scanning parameters and I do not see a difference.
@@nickdenniston4271 hello. Did you try the Optimize Threshold command first to determine the right settings for your image? Default settings work most of the time, but not everytime. You could also try deconvolving your images first before using the plugin.
@ ive deconvolved the images and i have also tried a variety of threshold setting indicated by the optimize threshold command. The best setting i found is a block size of 1um and a C value of 16. With these settings i am not getting as much noise and it minimizes the issue of having multiple mito being grouped into a single ROI. I am now worried that this C value is too high and i am missing information. Any thoughts?
@ I would think that a high C value is actually ideal for a deconvolved image.
Thanks for this amazing tutorial. I am trying to use this to do a ratiometric analysis using two channels. Is there a way to do it on a z-stack image and get the mean intensities in each of the channels?
Hi! Thanks for watching. If you want to do ratiometric measurements, maybe you can take a look at my other tutorial: th-cam.com/video/66NkiuTJtx4/w-d-xo.htmlsi=TTw2jk5A9ScTym1X
Great tutorial! May I ask what is the unit of the measured length. Is it in pixel or micron?
Hi there. The only unit that the plugin recognizes is micron. Measurements will be in microns.
Thank you one more time!
This is great, thanks a lot !!
Halo, if i want the measure the length of mitochondria which data should I use? total branch mitochondria percell or I choose the data from the total branch length per mito? thank you
@puspa_julistia1183, Per mito will give you measurements for each mitochondrial object.
Hi, i am trying to do mitoanalysis but i am getting an error, unrecognized command Sigma filter plus in line 194...Could you help me here please?
@@MrUnis hi ! I think it has something to do with the way you installed the plugin. Are you using Windows or Mac?
@@johanna.m.dela-cruz windows
probably yes, because in 3D Treshold optimimize .....I did not get radius and outlier remover
Hi, I tried several times to work with Mitochondrial analyzer function, Although I plugged in the mitochondrial analyzer but during using the 2D threshold function I am getting an error message like"marco or script not found".
Hi @sumansdiary8069. Are you using Fiji? Did you also install the required plugins: 3D ImageJ Suite,
ImageScience and IJPB-Plugins?
Hi !@@johanna.m.dela-cruz i am also facing the same problem. I am not able to get the jar files of these three : 3D ImageJ Suite, ImageScience and IJPB-Plugins.
@@promilalakra9886 hello. To enable these sites in Fiji, go to Help › Update…
Click Manage update sites,
Check ImageScience, etc.,
Click Close.
Click Apply changes
Restart Fiji.
Firs of all, great tutorial (most music and text ;o). If any one can help me, I had same trouble with the delimiter/units of the 3D Sphericity output. For a sphere, it is spected values from 0 to 1, but for same mitochondria I found values beetween 0.4 to 1.065. When I converted the data for comma, which I use it for a statistic program (Prisma Graph), I dont know if Im working with 1,065 or 1065,0; which for both cases is strange since the maximum expected value is 1. So, what Mitochondria Analyzer output parameters can I use, to recalculate the sphericity and to confirm these numbers?
Hi Guilherme. Thanks for your interest.
I have noticed this as well. I think that sphericity is calculated based on MorpholibJ's formula: "the ratio of V^2 by S^3, multiplied by 36*pi."
@@johanna.m.dela-cruz Thank you for the quick answear and for the equation. Now I will try to figerout how to "interpret" the number delimiter (point to comma)
if (canSigma) run("Sigma Filter Plus", "radius=" + sRadius + " use=2.0 minimum=0.2 outlier stack"..................why for thresholding in 3D, it does not recognize this command.
@@MrUnis Macros are not my strong suit, unfortunately. Did you use the macro recorder while using the plugin?
Hey I'm trying to measure the mitochondrial length in the same way. Although I can clearly see the difference in length between my control and test sample, still upon measurement the difference is not reflected. How do I manage that
Hello Arpita. Are you doing analysis on a per-cell or per-mito basis? If you want results for each mitochondrion in each image, you will need per-mito calculation to identify mitochondrial sub-populations. Per-cell basis should give normalized values rather than raw totals.
@@johanna.m.dela-cruz I did both. In the control vs test I can see the changes, but when measuring the mean length or length/mito i didn't see any changes
My another concern is also that if I change thresholding parameters in control or in test ami I not making the data biassed?
@@arpitadutta6737 The thresholding strategy of the plugin involves a local or adaptive thresholding algorithm that requires empirical determination of its settings. This means that finding optimal settings for thresholding values is largely based on your own observations. You will have to determine which thresholding algorithm to use based on your own judgement....Quantification will change based on the block size and C-values you use. S0, whatever method and parameters you choose will have to be applied to all your images (assuming they were acquired using the same settings).
@@johanna.m.dela-cruz heya. I tried to put adaptive threshold, changing the c value to 1 also didn't show any change, also I changed the block size into 0.25 from 1.125; that is the default. Still the thresholded image does not reciprocate to the original.
@@arpitadutta6737 Thresholding is always a challenge especially with complex images. I don’t know how your actual images look like, but you may want to try pre-processing them before using the plugin. Are your images 2D or are they z stacks? Some complex images might need deep learning algorithms for more accurate segmentation.
My ImageJ crashes when I try to do the thresholding. I can't proceed further 🙁 I have tried uninstalling and re-installing ImageJ. Does anyone know what's the issue?
Hi @BCBNarjis. How big are your images? What type are they (2D, 3D, 4D)? Are you using Windows or do you have a Mac?
@johanna.m.dela-cruz the 3D image is 108mb, and the 2D image is 4 mb. I tried both 3D and 2D threshold but they both crash the app without any prompt. I am on Windows.
Hey. I had replied to the comment but somehow it didn't post and I didn't realise.
Thanks for the quick response. I figured out that it was a memory issue. I allocated more memory and now it works fine. 😊
I had a 3D image, 108mb in size and I'm on Windows
Mitochondria Analyzer Manual PDF
Hi! Check the Description of my video for the link to the manual.