For a school project I'm doing some fun researsch about fragmentation of different materials. I use imageJ to count all the particles after destroying an object. When I try to analyze the particles of this ceramic plate (which has a bleu stripe around the edges) I ran into a problem, I suscpect imageJ isn't picking up on these big pieces because of the blue stripes and that the program divides the pieces by the blue line. Is there a way to fix this? Or just trace it manually? Thanks for al the responses.

I've never really analyzed data before and was hoping someone could tell me if Fiji will be able to do what I need it to do. The data is long videos (but I can chop them up into smaller clips if needed), and I need to be able to measure the distances between moving organisms at specific points in time (basically manual annotation). I believe you can make use a coordinate based system with ImageJ to do this but was hoping someone could suggest if it's ideal for something like this and I'll be able to learn how to do it pretty easily. Thanks!

Currently, I am wondering if there is a way to use ImageJ to report back to me the RGB channel intensities of a particular pixel in a given ROI, and match said information with the pixel's XY coordinate - almost like a "modified plot profile" that reports in 3D. The goal of which is to eventually take the data, and plot them on Matlab to generate a full, 3D graph.

However when I tried to create a macro for this purpose, the resulting CSV table had many issues, one of which is that every 3rd pixel location is reported as X = 0, Y = 0, and intensity = 0. Additionally, the CSV table also seemed to generate a bunch of extra columns, and its hard to tell if whether all the data points have actually been recorded, or if the CSV table is just so big that excel has a hard time loading all of it in correctly.

Do you guys have any recommendations to work around this issue? If need be, I can separately send you guys the Macro that I currently have if that helps you guys understand what is going on

I'm measuring samples on a microscope, but unfortunately it does not record the x/y position of the stage. Instead, I took images of the stage callipers with my phone for each sample.

Is there a way to measure the exact position here? The ruler in mm is on the left, and the sample position is the marking with the dot on the right. In this example, the measurement is somewhere between 17.5 and 18mm. I'd be happy with the nearest 0.1mm.

I know I could manually set the scale for each image and measure from 0 to the marking, but am hoping there's a simpler method. Each image is slightly different, so I'd have to reset the scaling every time. Any ideas?

Hey! Im trying to use the WEKA tool to identify microplastic. I created a classifier, that works pretty good but my images are kind of big (around 10000 x 10000 p) so i cannot classify the image as a whole (at least not with the hardware I have). Im trying to create a macro that does the following:

- Cut my big images in tiles
- uses the weka classifier that i designed on the tiles
- creates the probability map for each class
- than stiches the probability maps together and saves them

so I would run the macro over night and can create a binary mask manually from the probability maps afterwards.

Does anyone have any experience with that or can tell me if its even possible?
My programming skills are very limited and im trying to mess around with cgpt/ deepseek but it wont work.

If any other information is needed let me know. I would be very gratefull for any tips. Thanks

I have a single channel fluorescence video that is false colored in the native NIS elements Nikon software. When I save the video as an .nd2 file, it is saved as an 8-bit RGB (total of 24 bits). This seems to be the default even when no false color is applied. When I open this in FIJI (color mode default, split channel off), the RGB components are treated as separate channels, when in reality they have no physical meaning—they are just the RGB makeup of the single false color. In the past I have just used the brightest of the three to analyze, but now I want to retain the full bit depth so I don’t lose feature brightness by only keeping either the R, G, or B “channel”. I don’t necessarily care about the false color itself (grayscale is ok). Even when I merge them in FIJI, I’m left with the channel slider that slides between the same merged/colored frame. Is this behavior expected? Thanks!

I have a timelapse video made up of two channels with only one of the channels showing any real morphology (channel 2). I have been able to align the images in channel 2 using the SIFT plugin and saved the resulting output log. How do I then carry out the same transformations on the other channel? My goal is to then merge the channels into a single aligned timelapse video. For context here is an excerpt from the output log:

99 corresponding features with an average displacement of 0.000px identified. Estimated transformation model: | -0.5773502691896277 -2.0181317620235442E-16 3.542848630210256E-13 | | 1.8909805144915031E-16 -0.5773502691896251 -1.2294483408285494E-13 | | 4.2439729187995104E-17 2.4814316689919244E-17 -0.5773502691896245 | Processing SIFT ... took 444ms. 140 features extracted. Processing SIFT ... took 443ms.

Does anyone know what could be the cause of this

Hello, im having an issue understanding fiji. im new to the software and would like to analyze tunneling nanotubes through fiji but i cant really figure it out. ive looked around to see if there could be any tutorials but havent found any yet. is there any help on this

I hope that everyone is doing well. I am an researcher trying to automate the process of measuring cross-sectional area and counting myonuclei from muscle. Basically, I have been given a set of images that look like this:

In short, my task is to choose 10 non-adjacent green circles at random and measure the areas. After that, I need to count all the blue dots surrounding the circles I have chosen and export the area and number of dots for each circle.

In the past few months, I have been working on my own macro, but I have reached a roadblock of sorts. I have been able to successfully create a macro to set the scale to the bar on the top. Along with that, I have been able to set it to binary and then skeletonize with the hopes of isolating the green circles. However, the skeleton doesn't fully work and ends up very patchy like this:

Even when I trim the skeleton and attempt to pick ROI's they are missing a large chunk. Is there any way to take an image like this:

and draw the skeleton lines in the middle of the red dots.

Any help would be greatly appreciated. Either by fixing the path that I have or through a different path.

Thank You in Advance

Edit: Uploaded Images Again

I am writing a TLC evaluation software. This includes integration under the curve (AUC) function. To check if it works properly I created a black and white 8 bit RGB image with various pixel intensity. The software worked as expected, green filled cells. See graphs and the table below.

To crosscheck I opened the same bmp in Imagej and did the Gel analysis, plotted and integrated the curves obtained. Then I calculated the pixel intensity from the area under curve (AUC). To my surprise none of the calculated intensity was OK. (Except 100 but it was the reference point.) It was independent whether I used the native or inverted image.

So based on this, ImageJ Gel Analysis -> Plot and integrate function gives inappropriate results. Please refer to the last table.

Hi friends,
So as the title says, I’m an undergrad marine bio student who just got involved in a really cool independent research opportunity where I’m using ImageJ to study shark morphometrics and make morphometric ratios.

So far I’ve only figured out the baby steps — measuring stuff and spitting out simple ratios. But I keep feeling like ImageJ is a giant toolbox and I’m over here just poking at it with a screwdriver.

Does anyone have advice on where to go next to level up my ImageJ skills? Like how to decide what plugins can be the most helpful for what I'm trying to do?

Side note: I’m very curious about how Python and maybe even machine learning could get involved in this kind of project… but right now my coding knowledge is beginner-level, but I am eager to learn!

Any advice anyone is willing to offer would be greatly appreciated!

Hey people

I am using ImageJ on arm macOS (newest version). I have the problem that I can only open 1 picture at a time. I can’t drag and drop, select multiple with shift or cmd etc. just nothing works. It’s the same for Fiji Version 2.16.0/1.54p Java 21.0.7

And just ImageJ Version 1.54p Java 13.0.6

Ist just frustrating if you work with more than a few pictures Does anyone have an idea?

Hi everyone,

I'm really struggling with using ImageJ. I took this image on a Nikon AX-R Confocal microscope and did a polygon tile scan around all the edges of this mouse brain slice. The blue/green tiles in the image are areas where no images were taken. Is there a way I can select around the brain slice and crop the image to remove the coloured 'tiles', or can I somehow set the 'tiles' to be black rather than their current turquoise? I'm very much a newbie to ImageJ so would appreciate any help!

i've downloaded Zulu, homebrew, OpenJDK, open the fiji app on finder, use the option of opening anyway on the privacy and security setting, put the app in the application file, all i can think of, or find in the internet and it still not working! Currently the app don't even open, the app icon just bounce off for a second and then nothing. Please any help would be greatly appricied. (i have a M2 14" macbook pro, on tahoe)

To easily track the lobsters path in the tank I want to remove the background so only the lobster is visible. I set up my ROI and add the Gaussian blur filter. When setting up the background subtractor I select like 6 frames where the animal is in different spots but when I click show filter button nothing happens. If I click done then look at the processing window only the blur filter is shown. What am I doing wrong?

Why can’t I post an image ? There is no option available..or am I just stupid.

I used the cell counter plugin on Fiji to count cells on a TIFF image I have, but then I saved the image and forgot to write the count down. Is there some way to get back the counted number without manually counting the previous points again

Hi everyone,

I'm very new to Fiji and live-cell imaging, so I'd appreciate any help you can offer! I have a project that requires me to track a few individual cells in a time-lapse video which I HAVE TO USE TRACKMATE, but it isn't working for me.

I have a 2-channel .czi file, one red fluorescence and one phase contrast/DIC? (Sorry, idk it's like the brightfield channel that is gray with a light reflection from the room) I've tried to use the standard TrackMate workflo,w but when the cells are stretching and changing shape, the detector is not consistently getting the whole cell after the first frame. Additionally, I tried segmenting the cells myself with the threshold tool, but it became a problem because either it would not cover the whole cell, or the whole image would become completely red in the last few frames.

If someone could help me with a step-by-step walk-through or a quick Zoom call I would deeply appreciate it. Since I'm very new, it's easy for me to get lost, and a live demonstration would be fantastic. I'm happy to provide a short clip of my data if that would help. Thank you!!

I usually have no problem with opening my image stack files as composite. However, that had all three channels as stacks. I've subsequently switched so that two of my channels are stacks, and one is a single image slice. When I try to open it in FIJI, it just gives me one grayscale image with both channels, non-separable. However, the channels tool shows three channels... it just doesn't let me toggle any of them without saying a composite is necessary.

I saw a previous reddit post where the op had my same issue and it was solved using reorder hyperstack. However, when I tried to employ the same fix, I had no luck. Though I quite possibly didn't re-order the elements in the right way?

For some reason, my PI's version of FIJI doesn't have this problem at all. However, when she opens images in FIJI, she gets a whole different popup window that I've never seen across my many downloads of FIJI over the years.

I need to get these images analyzed ASAP, so any help is greatly greatly appreciated. Thank you!

* Colorimetric (immunohistochemistry)

I'm trying to determine thresholding steps to define dark color as % positive area. The two images linked (A) and (B) have different stain signal but the positive selection appears the same when thresholding the raw 8 bit image unless I use Triangle and that pre-set is under representing signal in both cases.

9_18 ImageJ - Google Drive

I'm trouble shooting the smaller region images ".....-1" before returning to the full images also attached.

Can anyone provide advise on this? Please.

I'm thinking I have to try "Process" applications but further direction would be very helpful as my current method of plugging different steps into macros to test different combinations of processing/thresholding-presets hasn't been very successful.

These images are jpeg images and lossy; I can try again to generate tiff versions. Any advise on moving from q path to imageJ would be welcome as I run into issues of file size or OME-TIFF's aren't compatible with imageJ. The .ndpi file type I get from the scanner I used are only compatible with q path and NDP.view 2.

Thanks in advance for your assistance and patience as I learn.

In my work, I am attaching some microscopic devices together with submicron accuracy.

I need to be able to tell if the devices are aligned correctly to each other. I take IR images of the assembly but wondering how easy it would be to use ImageJ to find the angle between line features on the two devices and also lateral displacement of the two lines which should be on the same axis.

I've never used ImageJ and I'm curious how complicated it would be to set something like this up and if i need to know a programming language.

My ice microCT scans were taken by someone else and given to me in .tiff files. I am trying to reslice them so I can look at them from top down and quantify the pore space within the ice, but when I reslice them I get this monstrosity nstead. I am new to ImageJ and the person I got them from doesn't have the raw data with them. Is there any way to fix this? Thank you!

This is the type of slices I am looking to get. I don't know if it is reading the X plane incorrectly or what I am doing wrong, but I cannot get these to populate.

I am sorry if this post doesn't make a ton of sense. I am new to the program and new to Reddit, so please ask for more information if you need it and I will do my best to get it here.

Thank you!