Confocal Microscopy - Science method
Confocal microscopy is an optical imaging technique used to increase optical resolution and contrast of a micrograph by using point illumination and a spatial pinhole to eliminate out-of-focus light in specimens that are thicker than the focal plane. It enables the reconstruction of three-dimensional structures from the obtained images
Questions related to Confocal Microscopy
I am using Glycerol 50% as my mounting media and not getting the required clarity. I have three tagged fluorophores. What can I use as readily available antifade reagent or how can I modify the mounting media to improve the clarity of my images.
I use this chambered culture well plates for immunofluorescent confocal microscopy: https://us.vwr.com/store/product/12361327/culturewelltm-removable-chambered-coverglass-electron-microscopy-sciences
I'm having issues removing the rubber gasket without breaking the attached slide. The slide is very fragile no matter what method I use to remove the gasket. I end up cracking the whole slide and rendering my sample useless.
Does anyone have any suggestions of a technique or a better brand of slides? I need ones that are for small infection volumes (30 ul).
Recently, I established a mutant strain of Drosophila with an APEX2 tag in its genome. Using this strain, I have successfully performed normal immunoelectron microscopy. However, it is difficult to detect the weak signal (it can be observed by confocal microscopy). Therefore, I would like to try the APEX2-Gold method. what points should I pay attention to when performing the APEX2-Gold method on animals? Also, can I keep the enhancement solution in stock? I would appreciate any tips anyone can give me.
Could you please recommend DAPI concentration for nuclei staining in mesenchymal stem cells during chondrogenic differentiation?
I have used 1 μg/ml DAPI for 6 min, then as usual PBS washing. but I got fuzzy colour and lots of unspecific binding (but still fuzzy).
If that matters - the brief protocol:
- PBS washing 3x
- permeabilization 0.2% Triton X-100/PBS, 4 min, PBS washing 3x
- blocking 0,2% BSA/PBS 1h, PBS washing 3x
- staining of collagen I, II and X (each with a pair of antibodies, with 3x washing in PBS in between steps).
I am overexpressing an endogeneous protein with c terminal myc tag in entamoeba. Overexpression has been confirmed by real time pcr. But when i am trying to visualize it in confocal microscopy with anti myc antibody i cannot see any staining. Primary antibody added is in ratio 1:100 and secondary antibody ratio is 1:300. Please suggest that what may be the problem.
I see varying sources describing this dye.
One place says it binds via its hydrophilic groups.
Another says its a lipophilic dye.
Which is it, how does it bind to internal membrane structures like mitochondrial and golgi membranes?
Hello, fellow researchers! I am trying to develop a project to track the growth and development of embryonic and larval killifish (F. grandis and F. heteroclitus) in varying environmental conditions. Confocal or fluorescent microscopy are methods I am thinking of using, but I am unfamiliar with either of these techniques. I am hoping someone with more experience could help give me the pros/cons of both or provide resources to previous research using either technique so I can familiarize myself with what is possible and the methodology. Thank you for any help you can provide!
In my imaging of mouse brains, there are fluorescent dots that appear in the image from confocal microscopy which are not any of the target cells in my experiment. I was wondering if using a secondary antibody from a donkey and a normal donkey serum in the blocking could potentially cause this?
Dear Fellow Researchers,
We are analyzing adaptive immune response in bladder cancer tissues using immunofluorescene stain and confocal microscopy.
Setting pinhole at AU=1 (perfectly confocal) results is very low signal. Increasing it allows to collect more signal but may yield some over-emission.
What pinhole is acceptable for tissue sections?
I am growing some large (1mm) organoids in hydrogel. I am planning to stain the organoids for 3 markers + nuclei and image them by confocal covering all 4 channels of UV, green, red and far red. I would like to use the UV channel for staining the nuclei. I am using a clearing agent (RapiClear) clearing the tissue before the imaging in order to be able to see deep inside the structures. The problem is that when I use DAPI, after clearing, the DAPI seems to fade and is not detectable anymore by confocal. Does anybody have experience with SYTOX blue from Thermofisher for staining the nuclei for confocal? Is it stable after clearing the tissue?
Thanks a lot in advance!
I am facing a very intriguing phenomenon. My immunofluorescence and confocal microscopy analyses showed a nucleus that is positive for MnSOD (which should be present only within mitochondria). Please, find attached representative images of cells stained for either nucleus or MnSOD positivity, along with the merged image.
Of note, it should be highlighted that the anti-SOD2 primary antiobody we used is extremely specific and that Western immunoblots showed only the specific band.
Did anyone else find MnSOD (SOD2) in the nucleus in some experimental or physiological condition?
Thanks a lot in advance.
Looking for some useful tips for fluorscence/confocal microscopy of plant root tissue.
I am mostly working with the root tissue of Arabidopsis thaliana where I take confocal/fluorscence images of plant root samples expressing proteins of interest with GFP/mcherry tags also often with Propidium Iodide/DAPI stains. Usually, I don't fix the tissue and take images as such and here are a few problems I face most of the time and I will be glad if you can advise me some suggestions to fix or minimize the following issues-
1.The amount of protein expression in various layers of root tissue is variable ranging from low to very high expression so in a Z-stack often I get highly staurated layers or if I adjust laser power/analog gain to minimize that oversaturation I end up losing signal where protein is less expressing. What should I do in such case?
2. how do you prepare live tissue for imaging, like if you use any specific buffer that does not change the physiological state of the plant tissue but helps in maintianing the GFP fluorescence for a longer duration?
3. Also, no matter how carefully I prepare the slide I never get the root tissue lying uniformly over the slide which means I get regions with good focus and in perfect plane wherease at some regions it gets blurry, out of the focus, how to avoid it?
4. I too have problem with DAPI staining of the root tissue as in most cases barely I see any nuclei but most often I end up with images showing DAPI at the cell borders just like Propidium Iodide.
Thanks a lot in advance for all the incoming beautiful suggestions.
I am using confocal microscopy to visualise membrane proteins of oleosomes (in any case proteins surrounding the oleosome) extracted from rapeseed.
The oleosomes have been stained with Nile red to detect the oil, and Fast Green FCF (0.013%) to detect the proteins.
The core oil is easily visualised but I am struggling to clearly visualise the proteins which would surround the oil droplet.
Do you have any advice on how to improve the visualisation? It may depend on the microscope settings I guess.
Thank you very much to anybody who could help me
I am doing immunocytochemistry staining for splenic B cells from WT and Knockout mice for one protein. By FACS analysis, it's clear that the signal for the knockout protein is nearly absent in Knockout cells, but when I do confocal microscopy experiment for the same cells, the signal seems to be the same in knockout as in WT. Does anyone have an explanation or can recommend what to do?
P.S: I use the same primary antibody for IF and ICC and No background for the secondary antibody in the control.
I would like to test by immunostaining that a specific membrane protein is present on my bacteria's outer membrane.
I cannot use FACS for this purpose, so I thought of confocal microscopy as an alternative. I would just fix the cells to keep their membrane intact.
I have not found yet any methods for this. Could you recommend one?
If you have any insight please share it with me!
For my project, I need to find colocalization of my retrograde tracer (tiny red retrobeads) with dopamine neurons in (TH immunolabeling, alexa fluor 488).
My problem is that retrobeads are so tiny and I only have few dots of them inside cell. Also finding very sparse red beads in red autofluorescence background in VTA is really difficult under confocal microscope. Does anyone know how can I solve the autofluorescence issue or potentiate my tiny red dots to be distinguishable in red background?
I fix my tissue in PFA 4% and thickness of my slices is 50um.
I'm using JC-1 to measure the mitochondrial membrane potential by laser confocal microscopy. I used 100uM of FCCP as a positive control. Surprisingly, it caused lower green to red ratio and was significantly lower than the untreated cells. My cells are cryopreserved mice oocyte. On the other hand, FCCP did work and caused an elevation in calcium using Fluo 3 AM and caused plasmalemmal membranes depolarization.
I need to mesuare the change in quantity and size of lysosomes in different experimental condition and I have this fluorescent dye that stains acidic compartments (LysoTracker Green DND-26) in live cells but It´s used for confocal microscopy So I need to know if someone has ever used in flow citometry with good results and if that´s the case: Could someone give me a protocol ?
I was thinking about VGlut2 (presynaptic) and PSD95 (post-synaptic) for the excitatory synapses, and VGAT (presynaptic) and Gephrin (post-synaptic) for the inhibitory ones.
Should I define as a synapse only the areas of colocalization between pre and post synaptic marker? Or should I consider also the isolate VGlut2/VGAT and PSD95/Gephrin ?
I am trying to analyse the state of the gut epithelium by analysing by confocal microscopy the state of some adhesion proteins (to note, Claudin and occluding); however, these proteins are not exclusive of epithelial cells, which is why I would like to add a marker specific for this subset. Could you please recommend me any marker/combination of markers that makes easier the identification of epithelial cells? Thank you in advance
I made a script to try to convert 4 colour nd2 files (not stacks) to run in the process> batch function of Fiji (see https://www.youtube.com/watch?v=fU104OU3Kk4 at 3:28) to create single tiffs, of all colours, merged. When I check the results in Fiji I get only the DAPI channel from the script below. However, if I run the macro in Fiji itself without the batch command, just on a single open nd2 file, I am left with the file that I want! So I am not sure why it is only saving the DAPI. Does the batch command save the earliest tiff that it sees and not let the macro finish? I am an inexperienced Fiji user so I am not sure what I am doing wrong.
title = getTitle();
one = "C1-" + title;
two = "C2-" + title;
three = "C3-" + title;
four = "C4-" + title;
run("Merge Channels...", "c1=["+one+"] c2=["+two+"] c3=["+three+"] c4=["+four+"] create");
We would like to use an enzyme tagged with eGFP and eGFP-tagged alpha synuclein at the same time to observe them under confocal microscopy. However, this will give wrong readings, so we would like to replace eGFP with mCherry. There is a research used the same way but without a detailed explanation.
I will add the link of this research below:
Dinter E, Saridaki T, Nippold M, Plum S, Diederichs L, Komnig D, Fensky L, May C, Marcus K, Voigt A, Schulz JB, Falkenburger BH. Rab7 induces clearance of α-synuclein aggregates. J Neurochem. 2016 Sep;138(5):758-74. doi: 10.1111/jnc.13712. Epub 2016 Aug 4. PMID: 27333324.
thanks in advance.
I want to quantify the cell area of bEnd.3 cells (mouse brain endothelial cells) using confocal microscopy by ImageJ. Most of the available protocols need to ignore neighbouring cells where cell-cell boundaries are in touch with each other. However, endothelial cells show continuous morphology. I will be really grateful if someone can help me with the protocol for cell area calculations using ImageJ.
I also want to quantify the distribution of tight junction proteins in cytoplasm vs. plasma membrane of these cells. If there is any way by which this quantification is possible using ImageJ then please help me.
Any help is appreciated.
Thank you in advance.
Can we use deionized water instead of mounting medium for confocal microscopy to preventing photobleaching? Since all the parts of the tissue does not mix well with thick mounting medium and therefore micro-air bubble develops.
We are going to upgrade our NIKON A1 confocal system with the incubator for long lasting live cell imaging - the incubator should contain atmosfere humidifier, CO2, temperature control etc. Please share your opinions on what system do you consider best and why? What to avoid ? What may be beneficial? The microscope will be used not only for live cell imaging so the system should be reconfigurable . We are looking for something from the top shelf but easy to maintain sterile environment and user friendly.
I am doing a mammalian oocytes maturation, followed by nuclear meiotic evaluation using confocal microscopy, I mount the oocyte over 90% glycerol in a double sided adhesive tape, and I am curious because I don't want to smash the oocytes under the cover slip. is there any recommendation for a good commercial wax cushion to be used in this case. (oocyte diameter is 100 micron).
I transfected HEK cells with fluorescent ABCG2 transporter (multidrug resistance transporter) , it has a GFP probe attached, and I am performing a live cell confocal microscopy experiment, where I would like to determine the kinetics of Mitoxantrone (chemotherapeutic, far red fluorescence) accumulation in the plasma membrane of the transfected GFP-positive cells. I am doing it by measuring far red fluorescence over time. For the information, mitoxantrone is the substrate of this transporter and ABCG2 is pumping it out of the cell, but when I apply an inhibitor to it, the accumulation rate increases, proving that the inhibitor works. That is how I am testing some of the compounds designed to be potential ABCG2 inhibitors.
Now that I hopefully explained the idea, I can get to the quantification part. I am confused about how many cells/fields of view to use to have a considerable number of sample for quantification. How many transfected cells should I take picture of? 50, 100? Is it for example 100 cells in 3 different experiments, or it is 100 cells per experiment, 3 times ?
As I have to take pictures in certain time point, I can only have one field of interest, where I usually have around 15 transfected cells. How can I collect 100 of them?
Or should I use regions of interest, few of them per one cell, around the membrane and measure that? I am really lost, so please those with similar experience, help!!!
Using confocal microscopy imaging technique is it possible to calculate the percent apoptotic index? In some research papers it is mentioned but up to what extent it is reliable and is it necessary to validate using FACS analysis?
- Isolated red blood cells (protocol in .docx is attached)
- Did confocal microscopy (stained RBCs with CellMask Red (photoiodide))
- Observed red blood cells (circled in blue) and also some cells with spiky surfaces (circled in green) (image as attached)
- What are those spiky cells?
- Is it normal for my red blood cell extracts to have them?
- If not, did something go wrong in my sample preparation, and how do I get rid of them?
If there are two different Nps prepared for two different drugs encapsulated in a single polymeric system(co-delivery) then can the uptake of both the NPs detected using confocal microscopy? In this case two types of fluorescent dyes needed ??
In another case, for cellular localization expt which dyes except Lysotracker can be used?
how to detect and counted early apoptotic, late apoptotic and necrotic cell population using confocal microscopy?
Is flow cytometry is mandatory to count the population or without using flow cytometry the cell polulations can be counted using confocal microscopy?
For image analysis, confocal microscopy is best of course.
I am interested in studying biofilm formation as a function of multiple variables, and would like to go beyond spectrophotometric/colorimetric/microplate readings and to analyze biofilm structures microscopically as well. Unfortunately, fluorescent microscopy (FM), confocal laser scanning microscopy (CLSM) and scanning electron microscope (SEM), tools that are routinely used by well-funded research groups for this purpose, are currently not at my disposal.
To this end, I am considering to utilize traditional staining (e.g., crystal violet) of these structures followed by simple light microscopy. May I ask for recommendations on objectively/quantitatively analyzing the images that can be generated this way (perhaps through a specific software)? What I have found so far are all for FM, CLSM and SEM images. Thank you!
I want to perform cellular uptake and localization study of my siRNA-loaded nanoparticles. For performing both the experiments which fluorophore-labeled siRNA, I should order.
Hi, I am currently working on the colocalisation of 2 proteins and I plan to work it out with confocal microscopy.
In the school I am working we have the leica LasX software with the co-localisation license but it is only available in that one computer connecting with the confocal mic itself. As there are always booking on that confocal mic and the computer is always in-use. I am looking for other softwares for the analysis of co-localisation. Best if I could draw an ROI, shows the pearson's and manders' correlation.
Currently I know ImageJ plug-in JACoP. Any other recommendations? And what's the difference with using the leica LasX co-localization analysis with other softwares such as Image J?
Is there any requirements on the software I use for co-localisation analysis if I am planning to publish my data?
Does anyone have any experience/reviews of the EVOS XL Core microscope?
I want to image one protein and a DAPI stain in a standard 96 well, clear plate like this https://ecatalog.corning.com/life-sciences/b2c/UK/en/Microplates/Assay-Microplates/96-Well-Microplates/Corning®-96-well-Clear-Polystyrene-Microplates/p/3799
(rather than an imaging plate with black wells). Would this work? Or would there be too much light scattering?
I would like to analyze a 1mm in thickness skin sample using confocal microscopy. The sample is too thick to be placed between a slide and coverslip then sealed by nail polish. The idea is to do z-staking and I don't want to cut the sample. Is there is an alternative way to tackle this issue? For example, can I mount the sample using some sort of adhesive tape?
I am working on amplifying a signal for confocal imaging using the Labelled Streptavidin/Avidin Biotin (LSAB) technique and ran into a surprising outcome: No staining. This is odd as I've employed this stain previously using the traditional indirect method and see decent specific signaling so I thought by amplifying the signal I'd get better, not worse, staining. Below I'll leave the basic outline as to my design (barring washes in between and counterstains).
- Incubate mouse anti-antigen primary antibody at 4 degree overnight (1:100; used at same concentration as in indirect method staining)
- Incubate biotinylated goat anti-mouse secondary at RT for 90 mins (1:150)
- Incubate Cy3-conjugated streptavidin for 90 mins (1:1000)
Any tips as to how to optimize this protocol and get it to function would be greatly appreciated!
As part of my thesis, I'm looking for gfp-tagged bacteria or other type of fluorescent-tagged bacteria already constructed in order to use confocal microscopy. I've already checked on ATTC but I only found a gfp-tagged pseudomonas. Is there another place (other devices or a laboratory) wich provide that type of bacteria ?
Thank you for the consideration.
I am working with FFPE sections of mice ileum and am trying to image changes in the mitochondrial ROS production between a treatment and control group. I'm not sure if there are any stains or antibodies that can specifically target mtROS once the tissue is fixed and I have seen another researcher propose using MitoSox Red on tissue prior to fixation to get an assessment of mitochondrial ROS in the live animal. Has anyone else tried this successfully?
We also dabbled using a pimonidazole hypoxyprobe as a measure of total ROS, however the staining in the ileum and colon differ for physiological reasons. If anyone has some ideas as to how to better optimize this stain or an alternative method of detection that works well with ileal tissue, please let me know!
Hi guys. I'm writing my paper for the first time.
. Representative images of glucose uptake were obtained of wavelengths at Excitation/Emission = 488/516 mm by using confocal microscopy
Is this sentence sound weird?
Or how can i correct this sentence ?
Thanks for nice answering .
Hi, I'm doing deconvolution to confocal images using Huygens Professional software, and I'm not sure whether I should decide the right signal-noise ratio (SNR) based on the image stack projected by summing the slices (SUM) or by projecting their maximum intensity (MIP). Despite the stack projection obtained by SUM looks better resolved than the original image, it seems to have more background. On the contrary, the projection obtained by MIP looks cleaner in terms of background, but some of the finest structures disappear compared to the original image (look at the attached images). The same is seen by comparing the intensity plots. Then, I wonder whether I should focus in change SNR to obtain better SUM or MIP projection.
Thanks a lot!
We have a Zeiss LSM 700 attached to an Axio Observer.Z1 microscope equipped with a T-PMT (photomultiplier for transmitted light). When I use the confocal microscope I often want a transmitted light (phase contrast, Ph3) image superimposed on the fluorescence light channel. This gives nice pictures as seen in the first attachment.
However at some point - using the same settings and same protocol - all images from the transmitted light channel has a wave pattern, this is a fault that I do not want in my images. See the second attachment. This problem persists and I have not been able to resolve the problem, do anyone know the cause of this? What I might have to adjust to fix it?
I'm having problems with the Recombinant Anti-MAP2 antibody from Abcam (Recombinant Anti-MAP2 antibody [EPR19691] (ab183830)) antibody on my frozen, 4%PFA perfused, mice brain tissue.
I was wondering if someone could elaborate better the note from the application part of the data sheet; Antigen retrieval: Heated citrate solution (10mM citrate PH 6.0 + 0.05% Tween-20).
To my knowledge AR is not used with frozen tissue, as it is to rough, and can damage the tissue, while also causing tissue detachment from the glass!?
Does someone have any experience with AR and frozen tissue?
I did try using 1/100 dilution, 0,2% triton X-100 and 4%PFA fixation as described in the images part of the datasheet: Immunohistochemical analysis of 4% paraformaldehyde-fixed, 0.2% Triton X-100 permeabilized frozen Mouse Cortex tissue labeling MAP2 with ab183830 at 1/100 dilution.
However I did not observed cytoplasmic staining of my neurons (image as uploaded for reference)!!
My protocol was the following:
4%PFA fixation, 15 min RT
10 min PBS+0.2% Triton x-100
Blocking solution (3%BSA, Without Triton x-100, 1h)
Anti-MAP2 1:100 dilution in 1%BSA (try both overnight and 2h RT)
AF-647 1:500, 30-45 min RT
3xPBS, 5 min
DAPI, 5 min
3x PBS, 5 min
dH2O, 5 min
Antifade and coverslip.
The antibody works excellent in IHC-P.
Thanks for the help,
I am trying to characterize the microstructure (localization of carbohydrates, fats, and proteins) of coconut milk powder using confocal microscopy. From the review of the available literature, I have found that fats and proteins can be non covalently labeled using fluorescent dyes. However, I am having trouble deciding the labeling protocol for the carbohydrate. Two common methods that I have seen in the papers are as follows:
1. Use antibodies or lectins conjugated with fluorophores to label carbohydrates
2. Covalent labeling of carbohydrates with FITC
If I use the first method, the signal from antibodies or lectins may interfere with the signal from protein present in the sample.
In the second method, the FITC (probe used for covalent labeling) also shows an affinity for proteins. So, I am afraid that the use of such a probe may also interfere with the protein signal (from the food sample). Is there any other probe available that I can use?
What would be the best way to label carbohydrates such that I can simultaneously image three components in the food mixture?
I have large, long lipid structures (several µm up to mm) that are present on a carbon coated 400mesh copper grid. My aim is to image these in the TEM.
Using confocal microscopy I can confirm that these structures are bound to the grid and withstand washing and blotting. After each step I check the grid in the confocal and still can see the signal from these structures.
However, a strange effect happens when I negatively stain them( with 2% UFo). In the EM, I can still bright traces (obviously areas without stain) at places where the tubes have been. But the actual tubes are (mostly) not there anymore. There are some small "breaks" in these bright traces where I can see something that has to be a leftover from the lipid structure, but it doesnt seem intact anymore.
I have attached images to better understand what I am talking about.
Has anyone ever seen such an effect and might know how to prevent this from happening?
Thanks everyone for any help/input.
I am planning long-term studies of tissue remodelling in vitro using light-sheet fluorescence microscopy. I am aware of confocal reflectance microscopy and second harmonic generation microscopy for imaging collagenous extracellular matrix without labelling, but I have seen only "home-made" systems for light-sheet reflectance microscopy. I would like to avoid immuno-labelling collagen, as this would not show collagen formed since the last antibody incubation.
Does anyone have experience in such applications using light-sheet? Would collagen's autofluorescence give a reliable enough signal? I see people are often looking for ways to eliminate this signal, so it seems promising. The quality of imaging doesn't have to be as good as SHG, as that will come later in the project.
I have cell images from Confocal Microscopy and want to do Nuclear Morphometric Analysis. Can anybody share a detailed protocol to do so?
Thanks in advance.
I did a Western Blot for a protein that should only be present in hematopoetic cells. The WB shows a very clear picture, present in k562(hematopoetic) and not present in HeLa and 293T. However when using the exact same antibody for IF on HeLa cells we do see clear signal from our protein of interest (see picture). Can anyone provide insights to how this is happening? in controls without primary/secondary antibody we see no signal.
100- 200 nm fluorescent beads are available from suppliers but only excited at UV (360 nm) or green and red range (i.e., >488 nm). Also. If the objective lens to be evaluated have (NA < 0.8), what is the appropriate bead size to evaluate the PSF from your experience? as 200 nm beads were quite challenging to use for measuring the PSF.
Hi! I am about to use AGR2 protein in my antibody display system. I want to check wherther protein can bind to cell surface displayed antibody or not. I need suggestions on how to fluorescently label (with red dye) AGR2 protein in order to view binding and also if someone could suggest efficient and cheap method to label protein that would be really great. please give me some insight and useful tips!
Has somebody seen this strange aggregation before?
I don't think they are agrobacteria since they are GFP specific (only excitable by 488 nm laser).
Any input is welcome.
Certain proteins phase separate into liquid droplets in solution. One can think of these as in vitro membraneless organelles. I would like to find how fluorescent labeled proteins partition inside the droplets using confocal microscopy. I do see intensity change between outside and inside but would like to get some quantitative results from this. I understand I would have to do some calibration using free dye. I am not very familiar with this process. Could someone explain/provide links to how this could be done? Attached is an example of a qualitative intensity profile across a droplet where the labeled protein was concentrated.
Whenever I read about confocal microscopy, it is used in conjunction with fluorescence. Wide-field microscopy also has out-of-focus information blurring the in-focus image. It seems that the laserbeam could as well be originating fom behind the sample. Why is it, that confocal microscopy is limited nonetheless to fluorescent applications?
I am looking for fixation protocols to non-adherent cells for confocal microscopy analysis. Also, I would like to know for how long I can keep the cells fixed (for example, I want to check if I can fix/prepare many plates during the week and take images from all of them only on Friday).
I have dozens of confocal images to process in Fiji. It is neither practical nor a productive use of my time to do them all out by hand, so I want to use a macro. However, I am fairly new to research; I only started a few months ago, so I don't have much experience writing macros in Fiji. What I'm doing is taking an image split into 3 channels, applying the grayscale LUT, and z-projecting to max intensity. Then I manually adjust brightness/contrast depending on the quality, merge channels, and stack to RGB. I also add a scale bar, but that I adjust manually i.e. 10 microns, 20 microns, etc.
Does anybody know if such a macro exists (i.e. in the Fiji user guide) or how to write such a macro? I have made one by using the record function, but that turned out to be inefficient because I had to change the file name every time I ran the macro. is there a way to get around this? Also, how would I incorporate into the macro the pauses when I have to make manual adjustments?
I have different titanium surfaces scanned using confocal scanning laser microscopy (CLSM) and need to analyze their roughness (Str, Std, Sdr, and Sds values are needed). thanks in advance for your precious time.
I do not have a TRITC filter or other laser set available other than 488nm.
I want to quantify/count neuronal cells, pretty well filled up with the expression signal of a protein. The images have been taken via confocal microscopy at 10x resolution. Which software/application/program is best and accurate to count the cells ? I don't want to count them manually!
Your suggestions can help me to analyse my data efficiently!
Does anyone know a simply way (i.e. using free image softwares) to quantify chromatin condensation? Cells were labeled with DAPI and analyzed by confocal microscopy.
In addition to clinical examination, dermoscopy and RCM help in diagnosing AHM. Which would be a better option between the two? Why?
I want to compare the surface roughness among different samples from confocal microscope (TCS-SP8, Leica), for example, to get the Rq or SA value from the surface.
Does anyone know the method to do this, or is there some software to do the calculation?
Thank you in advance.
I've used the NIKON A1 inverted confocal microscopy a couple of times and I noticed the red stain at 540/565 nm gets bleached within seconds. The objective I use is an oil 60x. Could anybody please let me know how this can be fixed? Massive thanks!
We are operating a Bio-Rad MRC-1024 LSCM on an OS2/Warp computer that is failing. I'd like to upgrade to a newer computer and more recent software, but Zeiss has discontinued the LaserSharp software that would run our LSCM, and they have no replacement. Has anyone found an alternative software to run this hardware? Alternatively (and assuming no objection from Zeiss), could we borrow the (now obsolete) software to install?
I am working on Symbiodinium cells (symbiotic dinoflagellates) and am trying to calculate the mitotic index. All the papers I have referred to use DAPI. I want to know if there are other stains or techniques that an use to distinguish between the doublets and just single cells. Is the resolution of DAPI good enough for me to see the mitotic spindle?
I am trying to culture mouse embryonic stem cells ( JM8A3; C57BL/6N ) on coverslips coated with 0.1% gelatin for confocal microscopy. The cells grow well but once I start washing the media with PBS, the cells detach from the coverslip. Anyone can suggest a way to fix that?
what is the best way to grow mESC for confocal studies?
Thank you in advance
I am analysing an immunofluorescence dataset taken by a confocal microscope in one z-plane. For each cell and channel, the mean fluorescence intensity in the nucleus and cytoplasm are reported. I was now wondering if the relationship between the mean fluorescence intensity and the concentration is linear.
- For instance, if a cell has 10 times as much nuclear mean fluorescence intensity in channel 2 than another cell, can I reliable say that the concentration of the antibody target protein in the nucleus is 10 times as high?
- Similarly, if a cell has 10 times more channel 2 mean fluorescence intensity in the nucleus than in the cytoplasm, can I reliably say that the concentration of the antibody target protein in the nucleus is 9 times higher than in the cytoplasm (assuming that ca. 10% of what is labelled as cytoplasm is actually golgi/ER/mitochondria/etc. and that these organelles do not contain the target protein at all)?
I'm working on a project in my university. Among other things, I'm required to count the number of astrocytes in the image stacks I've been given, as well as measure the length and thickness of their processes, using Fiji. However I'm having lots of trouble recognizing what might or might not be an astrocyte in each stack. I was told to create a composite image and a max intensity Z project image of GFAP stain (for astrocytes) + DAPI stain (for nuclei) then manually count based on colocalization of the stains, but I find that this leaves me with somewhat confusing images where I'm not sure if what I'm looking at is actually an astrocyte or not.
I've left a Max Intensity Z project image as an example. In this case there are many areas where I'm not sure if it's a full astrocyte or just a part of another one. And when identifying what I could more safely assume is an astrocyte (like the one on the upper right quadrant) I still can't see if there is a stained nucleus that belongs to that astrocyte. I can see there is a nucleus in the same zone, however, it appears to be much bigger than the astrocyte itself (I'd expect the nucleus to be "inside" the GFAP stain, like I've seen in other pictures). Would you for example count that as one astrocyte? And are there any tips to effectively identify all astrocytes in an image?
Thanks beforehand. I'm sorry if this comes off as confusing, but I'm really not sure how to work with this.