ImageJ - Science topic

I am okay to use open-ended software to do FEM or conversion to do it. I am working on fibre-reinforced composites so if I can get it to fibre level, it would be fun

Hi guys:

I want to quantify my nucleus morphology, the picture is taken by electron microscopy.

I have tried some methods, such as ImageJ, cellprofiler, but the effect is not good.

Could someone help me how to split the cell, nucleus, I want to know the area, perimeters, eccentricity and some other perimeters.

Thanks

Hey all,

Posted this on the imageJ forums, but maybe someone here can help...

I’m trying to analyze some angiogenesis pics, and have finally gotten Angiogenesis Analyzer to finally run, but now am unable to determine the right settings. Every setting I change gives me some version of this pic below:

Output:

Here is the original:

Here are my settings (but I've tried changing a few of the numbers with no luck of making it look better):

I’ve tried following along with the documentation, but nothing seems to impact the outcome much. can anyone provide some guidance?

I have a few images for which I will be calculating grey values. I will be using Image J. I know how to do it but there are several drops that need analysis and clicking. I already tried recording in Image J but I was wondering if there's a macro that exists that lets you select the oval ROI and automates the rest- creates a mask to calculate this grey values there. Clicking each drop get tedious and time consuming. Also curious what everyone uses for their analysis.

I am currently working on a project involving 3D image preprocessing and nuclear segmentation, and I would appreciate any guidance or suggestions you might have on preprocessing and segmentation for nucleus on tissue sample.

Specifically, my workflow involves:

Any help or clarification regarding this workflow would be invaluable to me.

Hi all,

I recently started using a new Olympus system which saves multiple image files as a .ets file with an associated with a 'master' .vsi file. If I understood correctly, the vsi file contains the blueprint of the datastructure in the .ets file.

After saving my image acquisition of an overnight experiment, the Cellsense software crashed. This resulted in the .ets file (containing 14 GBs data) being saved without its corresponding .vsi master file. Does someone have any experience extracting data from .ets files or working with Cellsense that could solve the problem.

I would greatly appreciate your help.

Merlijn

Hi All,

my protein of interest is a plasma membrane-bound protein, but it also has a cytosolic pool. I would like to measure its fluorescence signal specifically on the plasma membrane.

Could you kindly suggest any methods to achieve this? I appreciate your time and input.

How do you guys count your bacterial or yeast colonies? My fungus grows like yeast with distinct colony dots. I usually rely on the Fiji app(ImageJ) to manually tag and count them. What’s your go to method? 🧫 #colonycount #fungi

Hi,

We have performed an immunohistochemistry for AB in brain slices from APP mice and we would like to classify plaques in dense-core and diffuse plaques in an objective way. I have tried measuring the integrated density with ImageJ without success. Does anyone know how to do it?

Thanks!

I want to treat fibroblasts with 24h MG132 and then look a the amount of cell death. However, I want to correct for the amount of cells in total, because they also prolfirate in 24hour.

My supervisor suggested that after 24hour I stain my cells with HOECHTS and PI and then quantify with imageJ. The amount of dead cells can be calculated with:

x amount of HOECHTS positive cells/ x amount of PI positive cells.

However, HOECHTS also stains apoptotic cells. I do not know if this will interfere with my results. Do you think that after 24 hours most apoptotic cells are aleady dead anyway (or at least a part, normalizing the results)?

Or is this an unreliable method?

Hello everyone, I am a first-year PhD student and I aim to measure the calculated total cell fluorescence (CTFC) in my confocal images using the stain CellROX Green. As you can see, some parts of the cell have high signal intensity. I believe that extracting the fluorescence as it is will lead to inaccurate data so I wanted to ask what is the best way to pre-process the photos in ROI with high signal intensity/overexposure before quantitative extraction of the mean grey value? I considered eliminating any overexposed pixels using the Threshold feature in ImageJ and eliminate pixels with an intensity of 200 – 255. I can apply this approach to all the images in my data set and proceed to calculate the CTCF after. What do you think of this approach? Is there a better way to pre-process these photos?

Below are examples of my confocal images. Thank you in advance for any advice. Looking forward to what you have to say.

Hello,

I performed immunocytochemistry and captured images with confocal microscopy. I wanted to measure the fluorescence intensity of different experimental conditions, may i know how many number of cells i can evaluate through imagej from each experimental condition?

TIA

Dear All:

I want to quantify the fluorescence intensity of images using fluorescence microscope.

I would really appreciate it if someone could tell me how to use image j correctly.

I am trying to determine area of particles of cubic and octahedral shaped zeolite using ImageJ software. During thresholding, the software counts different number of particles in different operations. what should be appropriate way to consider the number of particles during thresholding?

I need assistance quantifying migrated PBMCs using a transwell assay. PBMCs (50 000 cells in 50 µl) were seeded in the upper chamber of a Corning HTS Transwell 96-well permeable support plate (insert pore size 8 µm) in serum-free RPMI 1640 medium with GlutaMAX. The lower chamber contained medium supplemented with or without the chemoattractant, 10% FBS, and the plate was incubated for 24 h. After incubation, the non-migrated cells were removed from the upper chamber. The migrated cells that adhered to the underside of the inserts were fixed with 4% paraformaldehyde for 30 min and stained with 0.2% Crystal Violet for 10 minutes, and rinsed with distilled water. The stained PBMCs were imaged to assess the number of cells that migrated through the inserts. Since PBMCs are semi-adherent, the non-adherent cells migrated to the bottom of the wells, where images were also taken for quantification. While I can manually count the Crystal Violet-stained adherent cells in ImageJ using the cell counter, counting the non-adherent cells will be more challenging due to their more concentrated distribution at the edges of the wells (see image), especially in the chemoattractant group where more migration is expected. Any assistance on how to quantify this in a quick and easy way would be greatly appreciated! Please note that some cells do adhere to the bottom of the wells, making it time-consuming to harvest each well of a 96-well plate and count the cell suspension using a Countess. I have also tried Crystal Violet staining in the bottom wells to measure the absorbance with a plate reader, but the results are inconsistent, as some cells are washed away during the process.

I was given neutron tomograms of plant roots. I don't want to analyze them manually, whether they are different or not. For example, counting the number of nodes myself would be problematic. I saw that there is a program ImageJ, but as far as I understand, you need to manually select pieces of roots (branching points, stem thickness and other parameters). Having these data sets, you can probably use some kind of statistical test? Does it make sense to train a neural network that will give me the number of branching points itself? I saw articles on computer vision, but I do not have a goal to invent something of my own. My goal is to use the most effective and fastest way to compare the topology and quantitative characteristics of two tomograms. It is also assumed that we will have about 8 tomograms per root in order to save time. Initially, I thought that if these are 3D images, then we can use some standard parameters for comparing two pictures, but 3D is not an option

I have fluorescent microscopy images in LIF format. There were 3 fluorescent dyes that I imaged with 3 channels. two of the three signals are detected in very similar areas of on my sample, they are difficult to tell apart. Basically I do not remember which image is in the second chanel and which is in the third. I tried looking at the OME metadata but there wasn't anything useful I could find. I would like to know which channel captured which wavelengths. Is there a simple way to get this information?

Hello!!

I am doing histopathological analysis of kidney sections with PAS stain. I want to quantify the staining intensity of the sections. can anyone explain what is the correct protocol for using the color deconvolution tool in ImageJ software?

Thanks in advance,

Sreyasi!

ImageJ application perceives cells as background. It counts the particles in the background. How can I change this? I change it from the file invert section but it didn't work.

Dear All:

I have to quantify the fluorescence intensity of cells that are fluorescent to two different markers, so i'm interested in finding out if this is possible.

Previously I have quantified fluorescence intensity using ImageJ but only with one fluorocrome, and in this case I'm wondering if it's possible for the program to discriminate only the areas that are yellow (im using green and red) corresponding to those cells that colocalize both markers.

I hope I was clear with my explanation.

Thank you very much!

I have images of film formed on the bottom of a 50mL beaker, and I have been trying to figure out the surface area. I could easily use features on imageJ to do that, but I want to know which area is covered by the film and which is not covered. I have explored the threshold feature, and I think either I am not doing it correctly or I am not very certain what I am doing is right. Is there anything I can do or process to follow to accurately know the covered area and determine the surface area based on that? I have attached images of the empty beaker and the beaker with the film. Thank you.

I am a beginner with ImageJ and would like to know how to quantify fluorescence images of platelets taken with a fluorescence microscope at 40x and 100x magnification. If you could provide detailed steps for the quantification process and explain what to pay attention to when exporting image files in Thunder, I would greatly appreciate it. Thank you.

Hello everyone,

I am working with mammary fat tumors from mice. My goal is to characterize tumor-infiltrating immune cells between different tx groups. I did immunofluorescence staining with primary antibody e.g., CD4 and CD8, then incubation with secondary antibody e.g., Alexa 594, and DNA staining with DAPI.

For image analysis I am using imageJ software. I would like to count CD8 positive cells in imageJ rather than manually counting but am struggling. I searched in the internet for any possible solutions but I did not find anything that could help me. Is there anyone who is experienced and can probably assist me with this please.

I want to study the defectivity of hexagonally packed cylinders. I prefer to use methods like Voronoi diagram analysis or Delaunay triangulation analysis for the defect analysis in the SEM and TEM images. what are the software I can use for this purpose?

I came across some papers but they use customized software for finding defects. Is there a way to use ImageJ for Delaunay triangulation analysis?

I want to analyze some images (nearly 1000) in a loop. I want to analyze HSV and RGB. I have masked those images in ImageJ, which is binary masked. I tried to explore them in R, but all the results came as NA. I also checked those images (some of them) separately in R to determine whether they were correctly masked, and the result was in matrix 0,0,0 1,1,1. But still, the result is NA. I used a chatbot to generate and analyze code. Can anyone suggest any codes and packages?

I want to analyze some images (nearly 1000) in a loop. I want to analyze HSV and RGB. I have masked those images in ImageJ, which is binary masked. I tried to explore them in R, but all the results came as NA. I also checked those images (some of them) separately in R to determine whether they were correctly masked, and the result was in matrix 0,0,0 1,1,1. But still, the result is NA. I used a chatbot to generate and analyze code. Can anyone suggest any codes and packages?

Hi, I have a data which consist of sliced tissue of olfactory system. I need to do intensity analyse by Imagej , but whole layers of the samples are not the same since the bottom part of the tissue has always less signals. What do you advise about analysing those data?

I am trying to analyze the size of agarose beads in a cell counter chip. The image was taken using brightfield so I am having a hard time processing the image well enough to where ImageJ can differentiate between the different droplets. It would be a lot easier if there was a machine learning algorithm or open source code that can help me differentiate between the different droplets and record the size of each droplet. I attached the images with the droplets that need to be processed.

I have taken fluorescence images of the control and treated sample(Immunofluorescence, tissue sample) at the same settings. So I need to measure the change in fluorescence intensity of the treated cells as compared to the cells in control

We have a demarcation and analysis protocol in ImageJ for images of C. elegans stained with Oil Red O. However, it does not seem to be the best way and we were unable to find an easy-to-execute protocol.

Hello everyone. I am in the middle of performing western blotting experiments measuring expression of p-S6(Ser235/236) protein in total cell lysate.

Originally, I was quantifying my data using ImageJ, and normalizing protein expression compared to the housekeeping gene on the same blot. (Ex: HSP60)

However, I was notified by my PI that she read a paper where the researchers compared expression of p-S6 to S6 protein on the same blot. It sounds like the researchers simply stripped the blot and re-probed with unphosphorylated protein since they are both the same size.

Which approach is the best way to normalize expression of phosphorylated protein on a single blot? I am considered that stripping and re-probing the same blot will cause loss of protein & unwanted retention of the phosphorylated protein.

How to count multilocular adipocytes on a slide with a white adipose tissue sample stained with H&E? I had done the count on the entire slide, but the articles put it in % of cells or adipocytes/mm². The total area of the slide is too large to be photographed under the microscope at 5X. And I can only be sure if it is multilocular with the 40x lens. So, I have no way of taking photographs and measuring them later in the ImageJ program. The articles do not detail how I do this measurement. The objective is to compare whether one group has more multilocular adipocytes than another: checking browning activity. Please if you can help, I would appreciate it

Hi, I'm doing my first lab internship and I have a little problem, I fixed some fibroblasts with 70% cold ethanol and then I treated with gentian violet to study their morphology. Now I need to count the cells, but there are many of them. Is it possible to solubilize the violet and treat with DAPI? so I could count them faster in ImageJ

lease help me to learn ImageJ software to analyze the immunoflourescence images. i need to analyze cortical neurons in the brain of stroke model of mice.

In my case, I want to measure the curvature of blue-marked particles in RGB color images, but the only possible way I've found to do it is by manually tracing the particles' perimeter and measuring the curvature with the Kappa plugin. As there are so many particles to measure, I think it would be much easier to just select them (i.e. with a threshold) and later measure them all automatically (if that is possible in anyway).

Thanks in advance!

Hello,

We conducted three stains for Nucleus, Outer membrane, and a particular phosphorylated protein expressed in a subset of cells. After capturing random images, our focus is on enumerating cells expressing this signal within the overall cell count. How we can do this in imageJ?

Best,

Dear all,

I am working with melanoma cryosections from minipigs. My aim is to characterize tumor-infiltrating immune cells between different age groups, mainly by immunohistochemistry. I do multiplex indirect immunofluorescence staining with primary antibody e.g., CD4 and CD8, then incubation with secondary antibody e.g., Alexa 488 and Alexa 555, and of course DNA staining with DAPI. For image acquisition I am using Leica SP5 confocal laser microscopy, then for image analysis I am using imageJ software. My question is how to quantify double positive signals using imageJ? Is there an easy way to do that? I searched in the internet for solutions but they got me very confused. Is there anyone who is experienced and can probably assist me with this please?

Thank you so much in advance!

Image analysis by ImageJ or Cellprofiler. Does anyone know of any protocols or tutorials that can be used to develop fluorescence and traditional microscopy image analysis research (histological slides) with ImageJ or Cellprofiler and would you have any tips for this?

Could you help me in the study of polyploidy using microscopy images?

Hi all,

I have the plot profile in imageJ and I saved the data (Distance_(microns) & Gray_Value) for the analysis. Could you please suggest to me how to do quantification with this?

Thank you

I have slides stained by mercuric Bromophenol stain, and I want to use imageJ to measure its intensity? How can I do that.

Besides what's the difference between H&E and H&E2 in imageJ?

Hello!

I am learning how to edit TEM images in Fiji (Image J).

I want to highlight some specific areas of my images using colors.

Can somebody help?

Thank you!

Dear Researchers,

Please advise me on calculating the homogeneity of two mixed powders with ImageJ by SEM images. Articles, guides, and real case studies are welcome. I want to automate the image analysis process as much as possible.

Thanks for all suggestions!

I had a macro that worked perfectly until I updated FIJI, now the “results” window will no longer open when after run(Analyze Particles…).

This is my macro, can anyone spot where the error is? Have tried asking ChapGTP and debugging line by line but I cant find anything.

dir1 = getDirectory("Please Select The Source (Input) Directory "); dir2 = getDirectory("Please Select The Destination (Output) Directory "); list = getFileList(dir1); setBatchMode(true); for (i=0; i<list.length; i++) { showProgress(i+1, list.length); filename = dir1 + list[i];

if (endsWith(filename, "tif")) { open(filename); setMinAndMax(-66, 321); run("Apply LUT"); run("8-bit"); setAutoThreshold("Otsu dark"); setThreshold(75, 255); //run("Threshold..."); run("Convert to Mask"); run("Fill Holes"); run("Set Scale...", "distance=4.4053 known=1 pixel=1 unit=um"); run ("Set Measurements...", " area perimeter shape descriptors feret's diameter Limit display add redirect=None decimal=3"); run ("Analyze Particles...", "size=5-Infinity pixel display exclude"); name = getTitle; index = lastIndexOf(name, "."); if (index!=-1) name = substring(name, 0, index); name = name + "(outlined)"; saveAs("JPEG", dir2+name); close(); } }

selectWindow(“Results”); excelname = “Resultsferet.xls”; saveAs(“Measurements”, dir2+excelname);

showMessage(“Task Completed”,“Inclusion counting is complete and the overlayed images and an Excel spreadsheet have been generated in the specified output folder”);

Hello,

I have imageJ image processing software and would like to calculate and plot the curvature in the beam using this software. I searched online and it suggests downloading the PhotoBend plugin. Can someone suggest any solution for determining the curvature of a beam using image processing software?

Hello,

I recently found the EzColocalization plugin for ImageJ and I would like to measure the colocalization of mitochondria and lysosomes in a group of oocytes. However, I do not know how to manually mark the area of the ooplasm in which colocalization is to be measured in the program. It always marks it automatically, but due to the presence of the cumulus cells, it is marked very inaccurate. I tried to mark the ooplasms manually, but the program does not respond or displays an error. Does anyone have experience with measuring colocalization in a group of oocytes?

Thank you for your advice

Linda

Which commands to use?

Hi All,

I am working on microglia dynamics in vivo with two photon microscope,but I cannot find a method to mesure microglia process length chang,I have tried imageJ plugin MTrackJ,but it looks work only on particles.Can anyone propose some methods to work on?

I have attached the microglia morpholoy,red-retractions,green-extensions.

Thanks in advance.

Hi all!

I have generated some Z-stacks of both cells and pancreatic islets staining for lipid droplets with Nile Red. I would like to quantify the number of, diameter and total area of the lipid droplets as i have seen in many papers. All the papers i have read so far just state that they have calculated these values in ImageJ but don't provide any details of how it is done. Any help would be very much appreciated.'

Hi there,

I am trying to assess colocalization (overlap) of two signals. The experiments were performed on FFPE human brain aged tissue samples, which means I have a lot of background after capturing my images (including the high amount of auto fluorescence).

Using ImageJ, I applied a filter to decrease the noise and then I performed background removal by subtracting the mean value of a background ROI, this for each channel. I did not use thresholding because all my samples had different gain values due to the variability of the auto fluorescence and background.

After this I am trying to obtain the Meanders' coefficient through the Coloc2 plugin on selected ROIs (for each cell body within), but I am not sure if I should use the M1 value or the tM1 value. Since I did the background correction through subtraction, and not threshold, I thought I should use M1. But I've been doing a bit more reading and I am just more confused.

Thank you for the guidance and help!

(PS: during the staining we already tried everything we could to reduce the auto fluorescence and unspecific background).

Hi everybody,

i am totally new to the programm Image J and i tried to use it to estimate my confluency in my cell culture. I found some descriptions but they do not fit for my picture at all... could anybody explain to me, what to do and how to use the programm on my cells? Please find a picture attached.

Thank you so much!

I would like to measure the area and diameter of thyroid follicles on a photomicrograph using image J (see attached image). Does anybody have a simple method for doing this given the large number of follicles in a single image?

What I am specifically hoping is possible.

1) Get an area/diameter for the entire follicle including the epithelium lining (in the high mag image this is the dotted black line)

2) Get an area/diameter of the colloid (blue dashed line and star region)

I would like to do this for the low magnification image attached that shows the large number of follicles separated by "interstitium" which is connective tissue and blood vessels.

In trainable weka segmentation in ImageJ/FIJI, the default classifier is the fast random forrest with 200 trees and 2 random features per node. Do I have to change the number of features if I am segmenting the image to more than 2 classes? Thanks

I'm encountering difficulties when it comes to quantifying the blue-stained cells in the image I've provided. These blue-stained elements represent cells, while the remaining particles are pores originating from the membrane of 24-well inserts. My goal is to eliminate these pores and isolate the cells. Is there a method available to remove the pores and exclusively highlight the cells in the image? Any assistance with this task would be highly valued.

Hi everyone

I'm facing a real problem when trying to export data results from imageJ (fiji) to excel to process it later.

The problem is that I have to change manually the dots (.) , commas (,) even when changing the properties in excel (from , to .) in order not count the numbers as thousands, (let's say I have 1,302 = one point three zero two) it count it as (1302 = one thousand three hundred and two) when I transfer to excel...

Lately I found a nice plugin (Localized copy...) that can change the numbers format locally in imageJ so it can be used easily by excel.

Unfortunately, this plugin has some bugs because it can only copy one line of the huge data that I have and only for one time (so I have to close and reopen the image again).

is there anyone that has faced this problem? Can anyone suggest me please another solutions??

Thanks in advance

Hello everybody,

I analyzed a picture in ImageJ, but I encountered a problem with the threshold. I tried to count cells with analyzed particles. Before starting, I adjusted the picture by changing the contrast and applying a Gaussian filter. Until yesterday, I did it exactly as described, but today the program displayed a message: 'the threshold may not be correct (255-255).'

Can someone help me with this issue? Has anyone with the same problem been able to resolve it?

I use imageJ to quatify protein but there are too many Subjective factors

How to measure the diameter of plaque

I am capturing fluorescent microscopy images of a developmental process with a high degree of variability. I need to compile these images into a "gallery" so that I can see many/all images at once to look for possible phenotypic differences between conditions, which then I can measure through FIJI. The most basic way I can do this is by arranging the images in a Powerpoint, but this method feels manual and has some limitations (explained below). I am hoping someone may know of a better way to compile image data for investigation/curation, maybe through FIJI, R, or a different dedicated program.

Limitations of Powerpoint:

1. Cannot track sample group or ID: When you upload an image into Powerpoint, there is no way to check its filename. So I am forced to upload one image at a time (and make a text box noting condition and ID), which is slow and unideal for batch processing images.

2. Re-formatting and arranging is manual: Re-sizing, cropping, and positioning images on Powerpoint is very manual and there is no record of changes (so making scale bars is hard). The effort compounded when the need arises to re-size or re-position the images later in the process (see #3).

3. Cannot re-order images dynamically based on values: Because the process I am studying is highly variable, it is necessary to compare images at the same percentile between conditions. Thus, I would like to re-order the images by measured values (so that I can compare min to min, mid to mid, max to max, etc.). This is also useful for selecting representative images for presentations/papers.

4. Interactive/Shiny Graph: To further aid in comparing images based on values, it would be nice to have an interactive/shiny scatterplot of the measured values such that when you hover over/select an image the corresponding datapoint in marked in the graph.

Partial Solutions:

A. Keynote: Solves #1 because uploaded images retain the filename information. However, the file format is limited to Macs which limits data sharing.

B. QuickFigures: This FIJI plugin solves #1 and #2, but it is still clunky to use and cannot re-order images.

C. Image Data Explorer: This R package or browser program solves #4 but can only view one image at a time. It also has the advantage of easy annotation of the images by entering values into the corresponding data spreadsheet/dataframe.

Does anyone know of a way/program that solves any of these limitations? Or does anyone have a different way of compiling image data?

area of infarct in percentage of total ventricle area. excluding the background

We've been looking at counting amyloid plaques in mouse hippocampus using 20x fluorescence images (antibody D5452). However when using either ImageJ or Matlab code, the fill holes or watershed options have thus far not worked to account for these dense core areas or partitioning plaques. I was wondering if anyone had further suggestions?

Hi everyone, I am currently working on lung fibrosis research on rat models of interstitial lung disease, I have problems with quantification of lung fibrosis on ImageJ. I used Masson trichrome staining on my research. I have read several papers with several different methods such as posted on ImageJ website (https://imagej.nih.gov/ij/docs/examples/stained-sections/index.html), color deconvolution (Ying Chen et al.) and other macro (Hadi AM, et al.; Kennedy DJ, et al.) but I don't think the published macro can accurately quantify fibrosis on my images.

Any suggestion of automated or semi automated methods on how to do it on Image J? Or maybe someone can point out studies with macro text who can be tried aside from the aforementioned study?

Thanks in advance.

Hi everyone,

so to quantify the reactivity of microglia, I would like to measure how much of the Iba1 signal is covered by CD68 by using IHC in mouse brain slices imaged with confocal microscopy at 40X magnification. Since the difference between samples is evident by eye already, I would like somehow to quantify that but I have more or less no clue on how to do it :( I was more thinking of setting as ROI the full area of the confocal images and not single cells. Is something like this possible at all actually? What would your suggestions be?

Thank you in advance for your help :)

Best Regards,

Marsela

Edit: here is an exemplary image. all my CD68 signal is located in the Iba1, but what I would like to measure is how much of the Iba1 is covered by CD68.

The available memory keeps changing even if no file is open. The files that I want to open are large files of around 7-10 GB

I used Olympus CellSens software to take staked pictures (.vsi). Open the file using ImageJ FIJI with Bioformat plugin but picture appeared more. purple so how can I see the original colors of HE stained as I saw it under microscope.

Currently, I am trying to determine the fiber diameter in ImageJ using DiameterJ Plugin. Although I created segmented images using the diameterJ plugin, after analysis DiameterJ does not generate an excel file containing radius calculations. I attempted multiple methods, including downloading the software from multiple locations. It still contains errors. Therefore, it would be helpful if anyone who has effectively utilized the diameterJ plugin in ImageJ could share that software. Or any recommendations?

Our lab is using ImageJ to process and compare IHC tumor sections. These sections are of varying areas and have varied signal strengths, and we want to quantifythis to determine if there is a meaningful difference between treatment and non-treatment groups (ex: CD8 cells). How does one account for the area differentials between samples? Would measuring total signal and dividing by total area be accurate? Thanks!

I am looking for good and preferably free software that will visualize a set of x, y, and z coordinates. I haven't had much luck with ImageJ or MATLAB. Specifically, I am trying to visualize 3D single-molecule localization microscopy data points. Any suggestions are greatly appreciated!

Hi,

I'm new to this forum and I have a question for you.

Here's my question: My goal is to quantify the fluorescence intensity.

Let me explain:

I am currently using zebrafish as an in vivo model to which I administer dye-labelled tumor cells. After the injection, at various times, I observe the fluorescence emitted.

Is there anyone who is able to help me and who maybe already knows how to quantify the fluorescence starting from the image taken under the microscope?

I've found several ways to use ImageJ, for example calculating the CTCF value but it doesn't seem to fit my purpose.

I also saw that maybe it is possible to quantify the fluorescence with the "CellProfiler" software, is this true? Would that be a simpler quantifying method than imagej?

Are there any other software or methods you can suggest to be able to quantify the fluorescence?

Thanks a lot for whoever will help me

Alessandro

hello everyone,

I would like to ask how we can analyse CD45 cell (should red) by using imagej program automatic way but by accurate method ( i want the counting of the cell in field or the %)? do you think the threshold method could use or accurate ??? or there any way to do this analysis ???

Thank you for your help in advance.

Thanks

Hello All,

I am in need of Alpha EaseFC software to quantitate Immunoblot protein bands. I know other options (such as ImageJ) are available to for this purpose, but I am specifically looking for Alpha easeFC software. I searched for it online but unfortunately was not succeed to find this. Any help will be highly appreciable.

Best,

I have these H&E slide images where I've been trying to do an a color threshold measurement on them to count the nuclei, white space, etc. which is pretty simple to do manually but as I try to create a macro to automate the process due to the large number of images it always fails. I follow the simple procedure to create a macro Plugins > macros > record to begin making the macro. Then for the color threshold I go image > adjust > color threshold > set parameters > macros > measure (command+M) to do the measurements which I have outlined the color threshold parameters below. Once I save the macro afterwards and try to run it on the same image I initially did it on, it fails to give me the same measurements as when I do it manually. I get a message saying "There is no images open in line 43 run ("Measure" <)> ;" after I see multiple windows with the H&E images open that are black and white. I am running ImageJ 1.53a on macOS Big Sur 11.6.7 and thank you for anyone that can help!

Measurement Values when performed manually

1 2764800 198.826 33 248

Color Threshold parameters

Hue: 0-255 [Pass is checked]

Saturation: 0-20 [Pass is checked]

Brightness: 0-255 [Pass is checked]

Threshold method: Default

Threshold color: Red

Color space: HSB

Dark background: Checked

Macro script

run("Color Threshold...");

// Color Thresholder 1.53a

// Autogenerated macro, single images only!

min=newArray(3);

max=newArray(3);

filter=newArray(3);

a=getTitle();

run("HSB Stack");

run("Convert Stack to Images");

selectWindow("Hue");

rename("0");

selectWindow("Saturation");

rename("1");

selectWindow("Brightness");

rename("2");

min[0]=0;

max[0]=255;

filter[0]="pass";

min[1]=0;

max[1]=20;

filter[1]="pass";

min[2]=0;

max[2]=255;

filter[2]="pass";

for (i=0;i<3;i++){

selectWindow(""+i);

setThreshold(min[i], max[i]);

run("Convert to Mask");

if (filter[i]=="stop") run("Invert");

}

imageCalculator("AND create", "0","1");

imageCalculator("AND create", "Result of 0","2");

for (i=0;i<3;i++){

selectWindow(""+i);

close();

}

selectWindow("Result of 0");

close();

selectWindow("Result of Result of 0");

rename(a);

// Colour Thresholding-------------

run("Close");

run("Measure");