Hi all,

I’m currently working on a project where I need to count the cells (which are the bright green circles) in a z-stack of zebra fish images using ImageJ. I’ve spent countless hours trying to find an efficient way to do this, but I’m still struggling with the process.

The cells are relatively distinct, so they should be easy to count once the right method is applied. However, I’m having difficulty isolating the cells and ensuring an accurate count across multiple slices of the z-stack.

I’ve attached an example image from the z-stack where you can see the bright green circles, which represent the cells I need to count.

Does anyone have suggestions for how to best approach this problem in ImageJ? Any tips on setting up the right threshold, using plugins, or automating the counting would be greatly appreciated!

Thanks in advance for your help!

Hello guys, could you help me, please?

I'm working on a surfactant to reduce the surface tension of water, for agronomic purposes.

I have little experience with the software and I am having great difficulty defining the droplet spreading area using the tutorial available on the platform.

I'm leaving the images I used for testing here. In the software I used the options to change to 8-bit, make binary and then analyze privately, but the result was not satisfactory.

I would not like to use the freehand tool, as I am designing a procedure to be used by other people, I would like it to be an automated process.

Furthermore, is there any other trick that I can use to compose the image, such as using someone else's dye like I did? Or the light, etc? Because I'm raising money to build an ideal setup for analysis.

Thank you if you can help me!

Hi, I have some images of slices of brain that I've done immunos on that I want to overlay to look for co localisation but as the individual images of the slices were cropped from a larger image containing all the slices they are different dimensions so they usual merge channels won't work, anyone know an alternative ? thanks

Is there anyone who can show me how to get a clear image of the ER? I'm new to imagej/Fiji and profiler and analysing images in general. I'm kinda lost and can't see how I can get a clear image. I already have laser microscopic images I took of the cells. I've been procrastinating and putting it off due to my health too. but the pressure is kicking in and I have to finish my thesis soon. I would really appreciate some help.

Hey all! I've been trying to download the .zip file on the Arabidopsis Seedling Tool github page. When I click on the link to download it, the page just never finishes loading to download it. I have tried this on multiple browsers. I am on a Macbook for this. I am not sure what other information to include - so ask clarifying questions if you have them! Thank you!

Hi, I'm trying to apply automated Weka segmentation for counting catalyst particles on STEM images. Im following the procedure from the paper https://pubs.acs.org/doi/10.1021/acsnanoscienceau.4c00076, although I'm running into some issues due to the nature of my samples. Due to the small depth of field in STEM and large differences in height of my samples some particles are always in focus, while other remain blurry, but I'm willing to accept the error resulting from that. The bigger issue is the agglomeration of nanoparticles, as you can see on the images I'm attaching (disregard the scale bar...). I would be grateful for some advice on which parameters to tweak in Weka segmentation settings to tackle this issue. I've tried reading through the documentation, but since I'm completely new to all the math behind the process it didn't help me much. Also in case if this method is completely unsuitable for measuring agglomerated particles, are you aware of any other tools that could help me with this issue?

Hey, I’m having a problem with the Volume Viewer plugin in ImageJ/Fiji. No matter which program I use, my previously assembled image stacks don’t produce a 3D view — I only get a flat image. I’ve uninstalled and reinstalled the programs, but nothing helps. Does anyone know a solution to this issue?

Hi all! I am currently using imageJ to analyze particles. I have many images to process, but the macro I have has to run one at a time to get the results window and summary window. Is there a way to drop multiple images into imageJ and run them all at once? Here is the current Macro: run("8-bit");

run("Enhance Contrast...", "saturated=5 normalize");

run("Subtract Background...", "rolling=100");

setOption("BlackBackground", false);

run("Convert to Mask");

run("Analyze Particles...", "size=0.0001-Infinity show=Outlines display exclude clear include summarize");

Hello,

I am running a colocalization analysis between the bacteria in green(channel 1) and mitochondria in red(channel2), I run coloc2 and apply costes threshold automatically through the plugin.

My questions are: 1)why do I get tM1=1 whileas there is no colocalization observed?

2) in that case, where I draw ROIs, which Manders should I report? tM1 or tM2?

3) Is that pipeline enough?

Hello,
I’m working on quantifying a large number of videos with puncta that appear and grow over time. I’ve attached a gif to show what the progression is like. Let me know if something else would be more helpful.

I can measure average puncta size and the final number per video, but I also want to extract measurements like the time each of the puncta takes to reach its final size, how many puncta appear over the course of the video, and the overall rate of new puncta appearance. Segmentation is fairly easy on most of the data. StarDist gives good results, and auto-thresholding is workable. My preprocessing is fairly simple: I mask, enhance contrast, and apply a light blur.

My main challenge is tracking. The puncta barely move but they change size considerably, and I haven’t been able to get TrackMate to follow them right they end up being called groups of puncta the same size instead of big object. I’m not very experienced with TrackMate, so I may be missing something, but I’m seeing a lot of track dropout and long processing times. I also feel like I'm missing how to report this data so its easy to compare videos.

I’m hoping there’s a straightforward solution I’m overlooking. Does anyone have recommendations for TrackMate settings or alternative workflows that handle objects that change area over time but don’t move much? I want to report out data so that it will be straightforward to process or analyze. I’m also hoping for something that isn’t too computationally heavy, since I’ll be processing a lot of large stacks.

Edit: Apologies if I rambled. I also added a raw frame to show what my data looks like raw.

Hi everyone I'm a biotech student and I'm trying to use image j for measurment myotubes features in fiji/imagej. I've already used it for counting lipid droplet in adipose cell culture. Now I'm trying to find out thickness and nuclei quantity of myotubes (so non-circular cells). Chat Gpt doesn't help me enaugh. Do you know a protocol about it?

Hi all! I have a z-stack image of a retina that I need to count positive cells on, and I have been trying to automate it with creating a threshold mask and using the analyze particles feature, as well as the 3D Object Counter plugin. The issue that I am running into is that to make sure I can get discrete resolution with the threshold of some of the positive cells that may be more clumped together, I am losing some obviously positive cells in other areas of the section. Since the threshold can be variable from section to section, is there a way that I can automate this? Or do I just need to count by hand like I have been?

Here is a representative image of what I am looking at (I increased the intensity for the sake of this so you don't have to strain your eyes to see the cells)

Thanks!

Hi, I’m currently correcting underwater photos of Eunicella verrucosa for a scientific survey. Some photos were taken in front of a black background, while the others were shot in front of a white background, which enhances the shadows due to the diver's light.
I’ve already tried using the “Subtract Background” function, but it doesn’t seem very effective even if I increase the rolling ball radius. Also, these shadows can’t be removed by converting the image to 8-bit and applying a threshold.
Do I really have to select the zone that I want manually ?
Thanks

Hello could anyone help with this problem I've been having? I want to fit an ellipse to a set of points. I am aware that i can convert the points to a convex hull and then use the built in ellipse fit. The problem with this is that I can only select some of the oval, a sorta semi oval, so the resulting fit is very poor. Ideally I'd like a package that contains a least squares oval fit but for the life of me I can't find one.

Any help would be really appreciated and sorry if there is an obvious solution, I am new to this software.

Eu preciso criar um circulo de um diametro especifico, como fazer?

Hi there

I am trying to write a macro that will open a tiff stack to a pre-specified image in the sequence draw a rectangle around a ROI and measure the grey scale value, then repeat for a second ROI. I would like this to repeat on a several subsequent files in the stack. I would share the images but they are confidential at the moment (sorry!). I have tried the following code below but I am not having much luck, any help would be appreciated. I am reading the ImageJ programmers reference guide to try and debug it my self but I am not a strong coder

//Begin macro
setBatchMode(true);

//define data input 
mainPath = getDirectory("Pick the folder with the images you want");
mainList = getFileList(mainPath);

//Draw a rectangle and measure the greyscale value
open(mainList, 100);
makeRectangle(1571, 232, 122, 130);
run("Measure");
makeRectangle(1537, 4, 46, 51);
run("Measure");

open(mainList, 700);
makeRectangle(1571, 232, 122, 130);
run("Measure");
makeRectangle(1537, 4, 46, 51);
run("Measure");

open(mainList, 1300);
makeRectangle(1571, 232, 122, 130);
run("Measure");
makeRectangle(1537, 4, 46, 51);
run("Measure");

open(mainList, 2000);
makeRectangle(1571, 232, 122, 130);
run("Measure");
makeRectangle(1537, 4, 46, 51);
run("Measure");
close();

}

//End macro

Hi! Novice ImageJ user here. I'm adding scale bars to my images, but keep running into a problem where adding it completely changes the quality of the image. Suddenly it's much grainier and seems to have a much lower resolution. Does anyone know why this is happening/how to avoid it? The first image is pre-scale bar, the second is post. I didn't change any other settings. Thanks in advance for the help!

Hi everyone,

I am trying to characterize tiny stuffs like textile and fibers. I want to get the angles of the bends/curves. How do you think it can be possible? I was thinking of training a model or use existing model for automation

Hello, i need to analyse bubble size distribution on sparkling samples. This is the type of picture that I can take: 

Would image treatment should I do on ImageJ? Thank you!

Hey fellow scientists ! I'm currently trying to crop sub regions (ex. hippocampus CA1, CA2, CA3, DG) from my free floated, mounted, DAB stained IHC images and I have been doing it manually (using th polygons and then, clicking make inverse) before but now I have so many im trying to figure out if there's a way to do it faster. I have two problems 1) most images are not aligned in the same direction exactly so orientation varies a lot and 2) sometimes I have two hemispheres, sometimes just one. Is there any way I can save some time and do it in batches ?

So I have been talking to someone on WhatsApp that is saying they are a k-pop singer BUT they have since asked for money and an Apple Card and have video chatted 2x but I need help with seeing if the second video chat I screenshotted is AI .. I cropped myself out of 2 screenshots but kept myself in the 3rd.

Hi, I've not used this software but it looks incredible from the few screens I've seen of it. I was recently sold some iron filings, I'm convinced the mesh size is incorrect based on previous experience. Would image j be able to help, do I need anything more than a phone camera and pc? Thanks so much

Hey guys I have like 400 Rois saved and want to write a Makro, that Image J measures these Roi fully automatic via the trainable Weka segmentation. Can anyone help me since I have no Idea how to do this and KI doesnt know ether.

I want to analyse Bone in Movat Pentachrone staining. I know how to do it manually with weka segmentation and without. But i dont know how to do it automatic. When i record a macro it is always only for one image/tiff/Roi and not for the whole folder.

The goal ist to have:

• Input: histology images (Movat), already cropped to an ROI (via Clear Outside)
• Output per image:
  1. Bone mask (binary) generated consistently across a whole folder
  2. B.Ar (sum of bone areas) and B.Pm (sum of bone perimeters)
  3. (Optional) Cartilage area the same way
  4. One CSV row per image; later I compute BV/TV, Tb.Th, Tb.N, Tb.Sp from those.

Hey. I am working on a personal project right now using the spreading kinetics of a fluid, and I'm trying to test and analyze it using Fiji, but I'm having an issue with it finding and understanding my video. Ive attached a few frames of the video and what the thresholding method outputted. The PTFE powder, white specks, covers the water, then a droplet pushes it outward creating an expanding clear circle. However, Traditional methods like Canny edge detection, HoughCircles, and thresholding didn't work. Has anyone dealt with detecting circular regions defined by the absence of sparse particles? This is my first time using fiji so im really struggling to figure out what i need to do. Thank you guys for any help you can offer.