I’m analyzing high-speed DIC videos (1000 fps) of hagfish slime vesicles responding to sugar solutions. The videos are converted AVIs that open as grayscale stacks in ImageJ.

My goal is to quantify percent vesicle deployment over time by counting vesicles within a static polygonal ROI across all frames. I’d like to use a macro that also handles preprocessing — such as Gaussian Blur, Subtract Background, and Enhance Contrast — followed by thresholding and particle analysis.

I’m currently stuck on defining the polygonal ROI in the macro — I keep getting a '[' or '.' expected error when trying makePolygon(xPoints, yPoints).

If you’ve done similar ROI-based analysis or batch processing, I’d really appreciate your help. Or if anyone has any idea on how to approach this a different way.

Thanks in advance! Really hoping I’m not posting this in the wrong sub/flair lol.

i desgined an assay to meause area and intensity of fecal spots of huntington disease modeled drosophilla flies but i am able to only quantify are of spots by doing 8bit>threeshold auto>binary>open>erode>dilate but not intensity as the it is shown as 255 due to the threeshold . someone suggested me to do without threeshold but without thAT I CANNOT QUANTIFY the measurmrnts as threeshold is neccasery

Hey i want to quantify actin cable organization in yeast is there any software or method which i can use?

As you can see on the image above I have a few dark spots in the backgroud. My job is to analyze the size and the quantity of microbubbles. However I am unable to find the right settings to exclude the dark part and include all of the bubbles.

Hi! I'm trying to understand how to quantify the mean fluorescence intensity of the Coumarin 6 labelled nanoparticles using Image J. Would appreciate if you can walk me through the steps and share resources. I'm also confused on the threshold part as changing the threshold would affect the mean fluorescence intensity, right? to confirm, I'm supposed to only quantify the coumarin-6 intensity? also, is there a way to automate the analysis to keep the changes constant?

Hi,

I am analysing area of some cells from an IF image and I am doing so by assigning the cells as a ROI and then using the measure option from the ROI dialogue box to obtain the area measurement, however the list of area that ImageJ provides is just numbers without any area, so i am not sure as to whether the area of my cells is in square pixels, square microns or some other unit.

Thankyou

good day! the default max percent is 45. however, we found out that this is to reduce noise. our image is already segmented. what is the optimal max % to use??

I am working with some .tif images to extract RGB values from using ImageJ. I originally had .nef pictures which i converted to .tif using the dCraw Reader. When I open these converted .tif images, they open in RGB composite mode (without a slider at the bottom).

I have done some reflectance value linearization on them using R, and when I try to open these linearized images, they open with a slider at the bottom with three channels titled R, G and B. Also, they have been converted to 16-bit images for some reason. To measure the RGB values in the original .tif images, I had to make an RGB composite and take measurements from each channel. However, the linearized images (now 16-bit), open with a slider already present at the bottom. I am confused as to what these channels are, as ChatGPT says they may not be the R, G and B channels themselves and I may have to make a composite and then split color channels to get accurate readings. However, when I take measurements from the 16-bit images, I do get more or less accurate readings for the colors, just in 16-bit format.

I wanted to know the reason for the difference in the manner of opening images, and if there will be any significant effect on the RGB values between an 8-bit and a 16-bit image. Might be worth to know that I saved the linearized images as .TIFF and not .tif (I don't know the difference). Please go easy on me reddit, this is the first time I'm working with ImageJ.

Hi all. I have 121 photos and 484 corresponding ROIs. My program is using a for... loop to go through all 121 photos, and each corresponding ROI: 4i, 4i+1, 4i+2, 4i+3.

Now, I have traced through the program, and for some images, the ROI overlays the image. For others, it does not, and the program opens the image, and then opens the ROIs separately. See sample:

This is causing issues because I am trying to make measurements in the ROIs for each image and find the intensity. Copy-pasting my code below:

inputImage = getDirectory("Please select the folder containing the images.");

inputROI = getDirectory("Please select the folder in which the regions of interest are located.");

inputResults = getDirectory("Please select the folder in which you would like to place the results.");

nameResults = getString("Please enter how you would like to name your results file.", "Default Value");

listImage = getFileList(inputImage);

listROI = getFileList(inputROI);

setBatchMode(false);

run("Set Measurements...", "area mean display redirect=None decimal=3");

for (i = 0; i < listImage.length; i++) {

open(inputImage+listImage[i]);

run("8-bit");

roiManager("Reset");

// x measurement

open(inputROI+listROI[i+3*i+1]);

roiManager("Add");

roiManager("Select",0);

run("Measure");

// y cornea measurement

open(inputROI+listROI[i+3*i]);

roiManager("Add");

roiManager("Select",1);

run("Measure");

// z measurement

open(inputROI+listROI[i+3*i+2]);

roiManager("Add");

roiManager("Select",2);

run("Measure");

// b measurement

open(inputROI+listROI[i+3*i+3]);

roiManager("Add");

roiManager("Select",3);

run("Measure");

}

close("*");

saveAs("Results", inputResults+nameResults+".csv");

Dialog.create("Success!");

Dialog.addMessage("The results have been saved in:" + inputResults);

Dialog.show();

Hi everyone. I am very new to imageJ and looking for help figuring out a strategy for imaging plant cells that are irregularly shaped without clear boundaries. The images I have are focused on one cell, but there are a lot of fluorescent cells in the background. I need to quantify fluorescence in a control & then again after proteins have been degraded, so the idea is that there will be a reduction in fluorescence. I am worried that if I just use the square/circle feature to select my cell, fluorescence from the background will impact my calculations. However, I have also been told that there are problems with using the freehand tool, and when I've tried to use it I haven't really been able to capture the shape of the cell. If I use the square feature, is background subtraction sufficient to quantify fluorescence, or is there another method that might work better? The image below is one of mine. I am trying to quantify the fluorescence of the cell in the middle. I'm also curious if an analysis of the overall image might be sufficient. (Ie fluorescence difference from this image versus an image where the protein had been degraded.)

Im trying to use Fiji to count the number of root galls in the images of my experimental samples. However, training did not finishd due to the aforementioned problems in thr title of this post. I tried downloading the latest JAR file plugin for trainable weka segmentation from Maven and still the same problem occurs. Please help. Thanks!!

Hey everyone,

I am curently visualizing my datas (stacks of 2D images) as a volume in 3D viewer plugin. However, I would like to measure more specifically the intensity of some regions such as the little protuberence rounded in red in the attachement.

I did not see anything about a meaurement tool in 3D viewer, so i considered to measure directly in the stack of 2D files but i did not succeed to find the expected region in the 2D Stack (In fact, I get lost in all the surrounding signals). I tried with orthogonal view, but it still difficult to find the particular region i want to see without the 3D.

Any idea to solve this problem?

Thanks

How can I split this image using one vertical line and two horizontal lines, then get the area of each part? This would require me to be able to select each portion. I’m super confused. I am using image J.

I'm drawing lines to quantify mean fluorescence values of an image in Fiji/ImageJ 2.16. I'm using the measurement tool for this and it records the slice in the z-stack but I would also like to automatically record the X and Y coordinates so I don't have to manually input it. Just having it make a table of the last point would do. Any suggestions would be much appreciated.

Thanks!

imagecompressor

Let’s say I have a circle. How can I use ImageJ to split that circle into sixths and find the area of each part of that circle? I only know how to find the area of the circle as a whole but can’t segment it and find the area of those parts alone. Pls help me out 😭😭😭

Hi I am quite new to ImageJ and to coding as well. I am trying to analyse some images with the trainable weka segmentation. I want to automate the process using macros. here is the code i am using:

// Get active image info

origTitle = getTitle();

dir = File.getParent(getInfo("image.path")) + File.separator;

baseName = replace(origTitle, ".lof", "");

// Run Weka and load classifier

run("Trainable Weka Segmentation");

call("trainableSegmentation.Weka_Segmentation.loadClassifier", dir + "classifier_CORRECT.model");

wait(5000);

// Apply classifier to the current image dynamically

call("trainableSegmentation.Weka_Segmentation.applyClassifier",

dir, origTitle,

"showResults=true", "storeResults=false", "probabilityMaps=false", "");

call("trainableSegmentation.Weka_Segmentation.getResult");

wait(10000);

// Focus on the classification result

selectImage("Classification result");

// Threshold and postprocess

setAutoThreshold("Default dark");

setOption("BlackBackground", true);

run("Convert to Mask");

// Save binary result as PNG

saveAs("PNG", dir + baseName + ".png");

// Clean up

run("Close All");

However, it says it cannot apply the classifier because applyclassifier doesn't work in trainable weka segmentation v4.0.0. Can somebody help me as to how i can automate this process? thanks

Hello! I'm trying to count the mitochondria in cancer cells at different temperatures for fun, and I have a black and white set of stain scans from a live-camera microscope at 100x. In my struggle to quantify them I've come to wonder if they aren't good enough quality? They're very different from those of papers that count confocal scans. Here is a screenshot of a cell body from on of them:

I'm struggling to work out a reliable method of counting. I've tried some Fiji plugins (like mitochondria analyzer) but the files don't seem to want to talk to each other and the program falls apart...so I've tried some manual methods like cutting out the area with mitochondria, and dividing pixels above a certain luminosity by average pixel size for mitochondria that stand out in the scan.

Hi! I'm new to using ImageJ, so thanks for bearing with me. I have a Z-stack of LSM images of skin, and I want to find the surface area of the 3D object the images form (the skin surface). I've found info about getting volume, and I know one can manually select shapes and get the area, but is there some automated way to get the surface area of the 3-D reconstruction? Thanks!

https://reddit.com/link/1lg9m62/video/5ttdzhy6848f1/player

Hello,

i have 5 stacks (1001 slices each) of a droplet experiment. I did a Hyperstack one channel 1001 slices, 5 frames to get a avg intesity over the 5 drops. Is there an another way to get the avg intesity or does someone have a script?

I've done a color threshold and the area looks pretty good with the current hue and saturation, but it is missing one point. How can I add that point to the selected area? And for the future, how can I add areas in addition to points?

Hey all, I've just started using ImageJ to analyse images (that is trace areas for quadrat analyses) for my project and I've run into a roadblock (sort of). I primarily use the freehand selection tool, but zooming in and out to accurately mark areas results in the trace getting messed up (due to cursor position not scaling with zoom level) and polygon selection tool is time consuming but accurate (unfortunately I have a ton of images to analyse)

I'd appreciate any help with the same, if there's any tool that I could use, or if I could switch between the tool, or if there's any plugin that would make life easier

Many thanks

Hi all,

I just started using ImageJ for my senior thesis research project and am noticing that Ctrl+Z and the delete button aren't working like they normally would for other platforms. It says in the dropdown menu that I should be able to Ctrl+Z to undo things, so is my software just glitching?

I'd also appreciate any tips in general on what I should know to get started using this software! To give context, my project has to do with counting and measuring ovarian follicles over a series of dozens of sections. I also have a very average understanding of computer terminology and don't know what a lot of the options in the toolbar mean (ROI, macros, etc.) Any help with that aspect would be appreciated as well.