Hello,

I would like to automate width measurements for segmented signals and I'm finding difficulty with the "bounding box" or "fit rectangle" resulting dimensions since I'm starting with diagonal signals. The longest dimension (length) matches my manual measures fine but the short dimension (width or height) doesn't match.

I tried duplicating a rotated-rectangle-ROI to make the signals horizontal but the binary image becomes grey scale due to resampling and I haven't found a way to keep my binary segmentation.

I followed this guide, https://www.youtube.com/watch?v=VJD36CY6aKE, to test "Local Thickness Analysis" and the average measure is at least closer to my 3 manual values.

selectImage("two segmented.tif");
run("Local Thickness (complete process)", "threshold=1");
roiManager("Select", 0);
run("Histogram", "bins=256 use x_min=0 x_max=4.47 y_max=Auto");
Table.showHistogramTable;

Is there any way to reduce the sampling to just 3-10 sites using "Local Thickness"? I have 70+ 'objects' per 1 image I am analyzing so 255 the histogram contains (per <2 microns) seems overkill.

Are there any other methods to accurately pull out this height value?

https://drive.google.com/drive/folders/1RvHwRA6MtwVaZ_yrMoP0S2qdpywrBgGY?usp=sharing I included some additional files at different processing stages but the simplest ones to follow are at the bottom right of the image bellow.

Thanks in advance for your time and help.

Perhaps this suggestion has already been made, in which case I offer my apologies.

I would like to suggest to ImageJ developers to move the File drown-down functions "close" and "close all" next to "quit". I understand people will probably tell me to just use keystrokes to avoid this problem, but I have used ImageJ for five years now and I still have accidentally closed work I meant to save. Yes yes, I should "get good" as the heartless will say, but it is also an easily rectified engineering control. I beseech you: these functions make no sense being next to "save" functions.

That is all. Thank you for your time.

EDIT: FijiCal is built from Fiji, because i find the workflow very painful. I havent packaged it for Linux or Mac yet because it needs testing before I got through those processes. I'm also working on a plugin only variant, but that's not a priority for me

I've been working on a small free utility called FijiCal, built around a deliberately narrow job: quickly calibrating microscopy images, taking measurements and producing accurate scale bars without needing to bring the image into a larger analysis workflow.

This isn't intended to replace ImageJ/Fiji. If I'm doing serious image analysis, that's still where I'm going. FijiCal exists for the other 90% of the time when I have a microscopy photograph and just want to calibrate it, measure something, add a scale bar and get on with my life.

I've now reached v0.9 “Mamanuca”, which is sufficiently mature that I can use it as part of my normal microscopy workflow without finding exciting new ways to break it.

The basic workflow is:

microscope image → FijiCal → calibrate against a known reference → measure / add scale bar → export

There's also now a Lightroom export plugin, since my own workflow starts with DSLR RAW files. I can develop the microscopy image in Lightroom and send it directly into FijiCal for calibration and annotation.

I'm releasing it free and would particularly appreciate feedback from people already familiar with ImageJ/Fiji. I'm interested in where a lightweight tool like this fits into your workflow, what it does badly, and all the horrifying edge cases I haven't discovered yet.

Current release: FijiCal v0.9 “Mamanuca”

Repo:
https://github.com/nighthunter226-but-real/FijiCal-Lite

Direct ZIP:
https://github.com/nighthunter226-but-real/FijiCal-Lite/releases/download/v0.9/FijiCal.Lite.0.9.-.Mamanuca.with.Lightroom.Bridge.zip

Bugs, criticism, workflow complaints and feature suggestions are very welcome.

Hi

I’ve been trying to reconstruct a neuron in SNT — I can do the dendrites no problem, but does anyone have any advice on how to reconstruct the soma in 3d instead of it being a sphere? When I do auto-trace with Area ROI, it creates the proper soma boundary, but when I go to the reconstruction viewer it still appears as a spherical mesh. Any tips appreciated!

Hi , I am an EE undergrad and I am struggling to create a map between the Ommatidia to its respective photoreceptors.

I cant find previous work doing this. Help we with the method please, it doesn't have to be this specific connectome. I want to know the resources and method to do it to get true vision for the Fly.

I'm a PhD molecular researcher, and like a lot of people here I've spent more time than I'd like to admit building ImageJ macros for Western blots.

The measurements were never the problem. ImageJ does the densitometry well. What kept breaking was everything around it: where the lane boxes go, how background gets handled, which exposure to use, and whether the next student in the lab does it the same way I did.

So I built that surrounding workflow as its own tool, Blotto (www.blotto.app).

You point it at your imager's export folder. It finds the lanes and bands, subtracts background, flags saturated bands, normalizes, and exports a PDF with the numbers plus every setting used to get them. Same math as you'd run in ImageJ. The difference is that it's the same workflow every time, for everyone in the lab, with a record of how each number was made.

If you've got a macro that works and you're happy maintaining it, that's a completely reasonable place to be.

This is for the lab where the macro lives on one person's laptop and leaves when they graduate.

It's a low cost paid tool with a free 14-day trial, no card.

I'd genuinely like to hear from people who know densitometry well: where it disagrees with your ImageJ numbers, and why.

I've tried using the 3D object counter alone and with adjusting threshold and binary mask. I can't seem to get values within range, does anyone have any recommendations?

Taste bud photos I'm working from

Hello everyone I want to start writing some macros for my analysis, I don't have a background in programing at all, but am eager to learn. What are some resources i can use to get started

is there a way to measure depth and size of a spine to classify spines? at least to my knowledge, the way to do this has yet to be found. Im still new to this so appreciate any help!

I am trying to count the cells in this image by threshold -> watershed -> analyse particles to count ROIs.

However, I cannot get the threshold to distinguish between the cells. Anyone know what to try?

Hi guys! I'm a beginner PhD microscopist wanting to find the best method to segment out and attach ROIs to the white nanoparticles in the catalyst clusters I am studying. Normal thresholding and "analyze particles" doesn't work here because the white nanoparticles often overlap - so a human can easily see two particles on top of one another, but I'm wondering if I can find a program that will do it for me.

I'm aware of StarDist, cellpose and other plugins but so far I've been unable to get a decent read (if you reckon you can get this to work, please tell me which parameters and thresholds to use!) I've attached my HAADF image of an example particle and the segmentation I've done by hand (using circles and free lasso) to give you guys a feel for what I want to achieve.

Here's a link hopefully to the png in question https://imgur.com/a/SP1stPi hopefully in better quality

Hi, I'm new to using ImageJ, and I was wondering if anyone has any suggestions for how I can get the vascular density from the attached image. I've tried a few programs, including Vessel Analysis, one made for a specific paper, and AngioTool (which didn't work due to Java issues). Any advice would be greatly appreciated. I also specifically want to calculate the density under the glass circular coverslip on the image.

Hi all !

So, I do live imaging of cells where I take four Z slices in two color channels every time point.

Due to what was likely a user error with the microscope, I have some movies where the first two Zs are inverted Is there a way to put them back in order in FIJI ?

I assume I would need to convert to a stack then back to a hyperstack with the right order ? But I couldn't find something that will let me decide the order of the Zs.

Thank you !

I have a 2D image and I’m unaware how to upload it or measure the area of my irregular ZOI. PLEASE HELP 😭

Edit: apologies for the stupid questions I’ve never image j b4

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.

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.

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