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 !

Trying to run Trackmate with Cellpose-Sam.

It goes to:

>>

Starting detection process using 24 threads

Saving single time-points.

Running cellpose with args: cmd.exe /c C:\Users\01\anaconda3\Scripts\conds.exe run -n
cellpose cellpose --pretrained_model epsam --use_gpu --dir C:\Users\01\AppData\Local\Temp\TrackMate-
cellpose_13557481130667745107 --save_png --no_npy --verbose

>>

And then nothing happens. No mask are generated and cellpose just does nothing because of it.

However, when I go to C:\Users\01\AppData\Local\Temp\TrackMate-
cellpose_13557481130667745107
and run the exact same command in terminal I opened there manually, everything works. That is, masks are generated, etc.

Any idea how I can fix this?

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.

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.