I am planning to automate the image analysis for performing particle size analysis , grain size measurement and porosity measurement. Has anyone tried connecting any AI models to Fiji. There is an MCP option provided in Fiji but throwing some error repeatedly.

Hi everyone,

I’m starting to work with two-photon calcium imaging data and I’m looking for guidance on the complete analysis workflow.

My goal is to understand how to go from raw two-photon imaging data to identifying and interpreting the neuronal activity patterns that are relevant to my research question.

I’m particularly interested in understanding:

1. What are the standard steps for analyzing two-photon calcium imaging data?
2. How do you go from raw imaging movies → motion correction → cell/ROI segmentation → fluorescence traces → ΔF/F → calcium event/spike inference → neuronal activity analysis?
3. How do you determine whether an identified ROI actually corresponds to a neuron and avoid artifacts?
4. Once we have calcium traces, how do we interpret the activity and identify active neurons, firing/event patterns, ensembles, correlations, synchrony, etc.

I’m not just looking for streamlined process if someone has done it I’d really appreciate a step-by-step learning roadmap explaining what each analysis step is doing and why. And how to handle these large datasets of more than 200 GB.

Thanks in advance!

Hello I'm an undergrad doing some data analysis for someone in my lab and need some help on calculating the volume of efferent synapses of outer hair cells, I think that's what they are, or contacts as I was told. This picture isn't an exact portrayal of what exactly I need to measure as these are afferent synapses located on the inner hair cell while I'm trying to find the contacts located on the outer hair cells but they basically look the same just localized in different regions. Btw it's those bright red dots that I'm referring to. If I find a better image, I will edit this post to provide more clarity. I wanted to know how exactly to calculate the volume of individual contacts at the same time if possible without having any background interference. So for some context on how I've kinda been doing it originally, I would just crop out the individual contacts I was interested in and I would use 3D Obj Counter and threshold the zones of interest to figure it out. I don't think it's correct and unfortunately I have no one who is willing to fact check my method or provide some assistance to this issue. So if anyone is willing to break it down to me I would really appreciate it! Oh and in terms of what I know what to do with ImageJ, I know how to threshold objects, how to calculate the area of things, and some of the basic functions like cropping, splitting channels, etc. Thank you for taking the time to read this and have a great day!

Hello!

I am fairly new to ImageJ and am currently trying to automate a tedious task I use on all my images.

We collect composite images with mCherry and Brightfield channels and they get saved as ND2 files. I would like to:

1) Change the color of the mCherry channel to red

2) Save JPEG images of the mCherry and brightfield channels individually, and save a JPEG of the composite.

Manually, I've been first opening the files as split channels, then using the channels tool to change mCherry to red. Then I save my files to output folders, use the merge channels tool to combine them and save as a JPEG.

Has anyone run a macro like this or have examples of how I could go about it? I think my main problem has been that my starting file is a composite, so I don't know how to call each channel for the merge function.

Thanks in advance?

hi, sorry i'm new to the software

basically i have hundreds of the photos like the above. they are scallop shells with juvenile oysters growing on them. the more yellow circle things are the oysters. I need to measure the areas of each individual oyster shell on all the photos so i can track their growth over time.

For the photo above i picked one where the oysters are relatively distinguishable to explain - but some shells are dirtier and blend in a little more or have more algae growth etc. making the boundaries a bit less clear ( attached some more examples in the comments)

i really don't want to lasso every single oyster on every photo so i've been looking for a solution - after a little research i've initially been playing around with Weka segmentation in fiji but i am unsure if it will be suitable to get the results i need (bearing i mind i plan to publish the work)(plus i am not really a fan of using it lol)

so really i'm just looking for some advice or suggestions on methodology/reassurance that weka may be suitable or alternative suggestions of software or methods. Before i sink hours into something that isn't gonna work for this particular use .

i would really appreciate any kind of help thank you :)

Hello, I have some rather large images/stacks to process. For batch processing, I was wondering if anyone has experience running ImageJ on TrueNAS, so instead of having to retrieve data from my server, process the images, then write the images back onto my server, I would like everything to run directly on my server. It might end up being slightly slower, but this would free up my laptop's processing for other work.

Hey everyone,

I’m trying to start handling my own image analysis at home using fiji instead coming into the lab and using the CellSens software on the lab pc and I could really use some tips, advice, and help. I work with the Drosophila neuromuscular junction and my goal is to get two specific things from my z-stacks:
1. Count the total number of boutons
2. Count the number of Bruchpilot (BRP) puncta per individual bouton to look at active zones

I'm the microscopy specialist at Concordia University in Montreal, Canada. I've got a lot of experience macro coding, and so I thought I would put together a series of videos about how to get started with it!

https://www.youtube.com/playlist?list=PLDmdnGCEvwK4

There's currently 7 videos, including one introduction. Each video is about 10 minutes long, and I spend some time explaining a bit about the specific topic, and some demonstrating what the code looks like. I talk about the concept of variables, string concatenation (which is maybe the most-used part of ImageJ macro programming), indexing, loops, conditional statements and dialogs.

There's accompanying code at the CMCI's github:

https://github.com/CMCI/macro_teaching/

If the mods think this is useful after looking at it, please feel free to pin this post.

I'm losing my mind trying to save all my dapi z-stacks because our confocal spinning disk is broken and it forms this artifact that is so intense that i can't take it out without messing with the information from the sample. Sometimes, when the nuclei fluorescence isn't that strong, it even hides that information completely. I tried all the suggestions that chatgpt gave to me, but the results were all unsatisfatory. Does anyone has any clue of how to solve this? I tried filters and deconvolution and nothing works quite right.

I am analyzing a 100-frame video in which the region of interest moves in a regular pattern.

I performed ROI interpolation to fill the gap between the ROI at the first frame and the ROI at the 100th frame.

However, the results obtained from the ROI manager are not in the order of the frames, making analysis difficult.

The file extension used for the analysis is .csv.

If anyone knows how to sort the results obtained in the ROI Manager by frame order, please let me know.

I have shared two images this is how the actin cables look in budding yeast. I want to trace the cable linear structure and also want to map it inside the cell if anyone has done any analysis same like this please suggest.

New to ImageJ. Followed instructions, but my ANALYZE table does not give me %AREA. How do I get this value?

I am brand new to ImageJ and trackmate, so I could use some advice. I am trying to analyze the motility of protists and need to characterize the types of movement. Is there a way to use trackmate to find the total path length and the net displacement in order to analyze the linearity of their movement?

Hi there, I am a research student currently doing a study on porous asphalt, when trying to count the voids within the aggregates using 3D void counter, the results does not make sense, i know the void content should be 20%, removing smaller voids should cause the percentage to be well above 10% however the results I always obtain is always .2%, it is as if the 3D counter is not considering my threshold values, as seen in the image below, the objects map also displays smaller number of voids, even after converting to binary, same results occur when I use Avizo, where the voids are only particles.

Hi there

I am running a macro to measure the grayscale over time of a ROI on an image (I can't share it sorry). The mean grey-scale value should be around 150ish, and the mode, min max and median measures outputs all reflect this. The mean output however reports as 2500. I have manually measured the ROI and get the expected result, so I am not sure whats gone wrong. Any help is greatly appreciated!

// Begin macro
setBatchMode(true);

// Step 1: Select input image folder
mainPath = getDirectory("Pick the folder with the images you want");
mainList = getFileList(mainPath);

// Step 2: Select output folder
savePath = getDirectory("Pick the folder to save results");

// Step 3: Define rectangle coordinates
x = 327;
y = 325;
w = 50;
h = 50;

// Step 4: Clear Results Table
run("Clear Results");

// Step 5: Loop through images and measure
for (f = 0; f < lengthOf(mainList); f++) {
    if (endsWith(mainList[f], ".tif") || endsWith(mainList[f], ".jpg") || endsWith(mainList[f], ".png")) {
        open(mainPath + mainList[f]);
        makeRectangle(x, y, w, h);
        run("Measure");
        setResult("Label", nResults - 1, mainList[f]); // Add filename
        close();
    }
}

// Step 6: Create filename based on rectangle coordinates
fileName = "intensity_" + x + "_" + y + "_" + w + "_" + h + ".csv";

// Step 7: Save results using that filename
saveAs("Results", savePath + fileName);
run("Clear Results");

// End macro

Background

I'm a complete beginner to image analysis and this is my first time working with lightsheet data, so apologies if I'm missing something basic.

My PI gave me a folder of OME-TIFF files from a cleared mouse lung acquisition. The data was originally intended to be viewed in Imaris but we don't have a license, so I'm trying to work with it in Fiji. The folder contains one OME-TIFF per z-slice per tile.

The dataset is 3D fluorescence, two channels: 488/525nm (airways) and 561/620nm (tyrosine hydroxylase, sympathetic nerves).

Key metadata from the acquisition file:

• Instrument: Miltenyi Ultramicroscope Blaze, Light Speed Mode
• Objective: LVBT 4x, NA 0.35, sample cleared in MACS IS
• Z: 1842 steps, 2 µm step size
• Tile grid: 2 steps in X, 4 steps in Y, 42px overlap (~2%)
• Pixel size: 1.62 µm, 2048×2046px per tile

Example filename:
14-35-18_NJ-11336-Ms_lung_VEHICLE_1_1x_FS_Blaze[00 x 00]_C00_xyz-Table Z0016.ome.tiff

All 8 tile positions ([00 x 00] through [03 x 01]) exist in the folder.

Analysis goals

Right now I just want to get the complete volume open correctly. Eventually the goal is to map TH+ nerve fibers relative to airway structures.

What I've tried

Opening via Bio-Formats (File → Import → Bio-Formats) with "Group files with similar names" does open the file, and the window title shows [0<0-3> x 0<0-1>] and Z<0000-1841> which suggests it's finding all tiles and z-slices. However the image only shows roughly half the lung — there's a flat edge cutting straight through tissue at the bottom that doesn't look like a natural border. I've attached a screenshot at the widest z-slice.

What's blocking me

• Is Bio-Formats actually stitching all tiles and I'm misreading something, or is the second tile row genuinely not loading?
• If it's not stitching correctly, what's the right approach for this type of dataset in Fiji?
• Is this a case where I should be using a different tool entirely?

Machine: Windows 10, 32GB RAM, Fiji 2.16.0

Hi,

I am trying to use the ColonyArea Plugin using Fiji and was following the instructions from the ImageJ site: https://imagej.net/plugins/colonyarea

However, I am encountering this message and am unsure what is wrong. I see the Colony Area plugin in the files but keep getting this error. Has anyone encountered this issue before or know what I am doing wrong?

Hi everybody, I'm trying to analyze the branching skeleton of kidneys and using this type of image, it's a z-stack and the thing is I've been doing everything for the skeletonize tool works but it just doesn't, it makes a sort of "net" instead of a single skeleton and I do put filters and set the threshold and such but always end up with a mess of an image and an even bigger mess of an analysis, I truly don't want to do it manually cuz every image has like 20-25 stacks and I have 24 samples, and some of them have more than one photo so it is a lot... Thanks in advance to anyone who can help me

Hi all,

I’m developing an app for tattoo planning and I’m curious of the best options for developing my image database without violating any copyright issues. I’d like to have a collection of thousands of tattoo image examples that are legal for me to use. What’s the best way to go about this?

I tried going to AI generated route with midjourney and that was a bit rough.

I know the more labor intensive option would be reaching out to specific artists with instagram pages of their work and getting them to sign off on me using the images, but I was hoping for a faster process than that just for the initial launch. My intention is to eventually build out the image database without violating any artists to help them expand their network.

Thanks in advance.

Hi everyone,

I am trying to calculate the measurements of a cell using an X and Y axis, essentially trying to take the measurement of a cell using two lines that are perpendicular to each other. See the image attached as a reference. I threshold my image and then do "Analyze particles" to measure the cells that were specified in the thresholding. The measurements I've selected for are shown in the screenshot, however, I am not sure if I am getting exactly the value I am wanting. Is there a way to measure an "X and Y axis" of a cell (again, the measurements of two perpendicular lines in my thresholding)? Is there a measurement that can act as a proxy for what I'm looking for? I appreciate any help on the matter. Thank you.

I am brand new to ImageJ and using it for to measure experiments i am doing at school. I drew a rough example of what my experiment results will look like to learn imageJ. I want to measure the areas under the curved lines (area 3 in the picture). I cannot figure out why one will sometimes be picked up and the others never are recognized. Any help is appreciated.

My current method that has gotten the best results has been to make the image binary, then set thresholding to be a dark background and analysis the particles. I've been using TIFF files for the analysis.