Fluorescence Imaging - Science method
Fluorescence Imaging is a fluorescent labelling and staining, when combined with an appropriate imaging instrument, is a sensitive and quantitative method that is widely used in molecular biology and biochemistry laboratories for a variety of experimental, analytical, and quality control applications. Multicolor fluorescence detection allows the detection and resolution of multiple targets using fluorescent labels that can be spectrally resolved, thereby permitting the detection as well as the quantification of proteins and nucleic acids.
Questions related to Fluorescence Imaging
I have developed a hydrogel based on silk, and I need to do some fluorescent imaging of the encapsulated cells. When I stain the cells, for example, using CalcinAM, the background is huge and does not allow good-quality images, and I think it is because of the inherent autofluorescence of silk fibroin. Do you have a way to circumvent this?
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?
I will locate a newly induced human protein in yeast to test its location correctly. The original position of this protein in human cells is ER. So I decided to add a fuse mCherry to it, then use fluorescence imaging to detect if it can locate at ER in yeast.
The question is where should I put it? N-terminal or C-terminal? Should I use some flexible linker between them?
In the fluorescence imaging, I am getting fluorescence at 800 nm instead of 690 nm. What could be the reasons behind this??
I'd like to quantify my fluorescent images. I want to compare 2 different regions, one is bigger in size and the other smaller. and I measure the fluorescent intensity mean in these two regions. Should I divide the fluorescent intensity mean with area of region?
I am trying to prepare Symbiodinium cultures for microscopy (fluorescence microscopy and potentially confocal). I don't know if i have to fixing my cells in glycolaldehyde (GA) or if i have to fix at all, because I'm trying to capture the chlorophyll fluorescence from my sample, and if I fix it maybe it will die and not be fluorescence anymore.
Could someone help me?
Thank you very much
I have images stained with different immunofluorescent markers (DAPI + many different markers) and captured images. However I recently discovered when I went to Metamorph to compare that the exposure times were different (300 vs 350ms). I know that in an ideal world, all the settings would have remained the same, however I am trying to salvage this data as re-doing the experiment is not an option at this time.
I have read the Filkins paper attached here talking about normalizing exposure times (Dark pixel intensity determination and its applications in normalizing different exposure time and autofluorescence removal) but my question is - how do I actually go about applying these changes to the images in question? Is there a journal I run it through?
I purchased the Cyto-ID autophagy detection kit from Enzo and would like to use our lab's fluorescence microscope to image the autophagosome punctate. However, the product manual says the preferred magnification is 60x and the best our microscope has is 40x.
Has anyone tried an alternative magnification with this kit and observed the punctate successfully? Thank you.
I am new to culturing cells in a 96-well plate. I am using BALB/c 3T3 cells and culturing them in a black plate with transparent bottom for fluorescence imaging. The cells remain distributed throughout the well after treatment with some compound. But I find that after staining, the cells will gather at the corners of the well. In the beginning I thought it was because I swirl the plate slightly every PBS washing. But after trying not to swirl, the same problem occurred. I also dispense liquid very gently and along the wall of the well. The second photo below is before staining and the first photo is after staining (the cells have gathered on the corner of the well in a ring like manner, leaving an empty space at the middle of the well). If you have any suggestions, please let me know. Thanks 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.
- 461.52 KBSHSy5y ELF SOD1 e SOD2 5 e 10 gg 21.12.2016_Controllo 5 gg SOD2 zoom 2 1 Series011_z0.tif
- 208.56 KBSHSy5y ELF SOD1 e SOD2 5 e 10 gg 21.12.2016_Controllo 5 gg SOD2 zoom 2 1 Series011_z0_ch00.tif
- 270.31 KBSHSy5y ELF SOD1 e SOD2 5 e 10 gg 21.12.2016_Controllo 5 gg SOD2 zoom 2 1 Series011_z0_ch01.tif
Can the rolling ball radius tool be used alone to reduce background in fluorescent images? Why or why not? If not, what are some additional methods to further remove background after rolling ball has been applied?
We're looking for the best system to test tumour targeted T cells against a cancer cell line. We've been using an Incucyte system that gives a fluorescent image based time course of tumour cell death. We're interested in what other systems, image based or otherwise, are currently available and reccomended.
I want to quantify fluorescence images from IHC experiments using Adobe Photoshop, but unsure of how to do so. If anyone has a detailed protocol or could direct me to one, I would be grateful for that.
We are studying biofilm formation on differently coated 15mm diameter uPVC discs - these are opaque and may be white or black in colour depending on coating. The current setup includes forming biofilms on discs, rinsing to remove non-adherent cells, fixing each disc to a glass slide and then adding a drop of LIVE/DEAD BacLight mixture (Invitrogen L7012) to the centre of the disc. After incubation, the excess stain is rinsed off, a coverslip is placed on top of the disc and the edges of the coverslip fixed to the underlying slide with nail polish. After drying, the whole piece is placed in the inverted microscope coverslip side down. When imaging it is possible to see a hue of green or red depending on filter but not to resolve cells, and it is unclear whether this is just autofluorescence of the disc or coating. The opacity of the discs seems to block much of the light.
Thank you for your assistance and advice.
I'm currently working on my bachelor's thesis about thermic effects in FRAP(fluorescence recovery after photobleaching)-experiments when using a confocal laser scanning microscope (CLSM).
Right now I'm trying to figure out what causes these increases in intensity around the bleach-spot that starts to shows up when doing FRAP at a certain depth. For all my measurements I set the height as 0 μm where the excitation with the laser caused the highest intensity. The increases in intensity started showing up at a height of 20 μm and persisted up until 300 μm (I went this deep into the sample out of interest). My samples were solutions of rhodamine B in glycerol and sulforhodamine B in glycerol (both with a concentration of roughly 1 g/L).
My research so far didnt give me any possible explanation for this phenomenon. I'd be glad if someone in here could either give an explanation or provide me with literature on this subject. I will add 2 pictures so you know what im talking about if my explanation wasnt clear enough. If some vital information is missing, I'll provide it down below.
Until then and thanks in advance,
I'm trying to stain living cells, but always in the fluorescence image I find that the size of the dye representing an object is larger than the original object in the bright field (as if the dye was on the outside of the object as well as inside). any suggestions? All I can think of now is that it's either insufficient washing or the dye concentration is too high.
I'm working on getting replicates of an ICC protocol and imaging+counting cells for analysis. I've noticed that, between replicates, there are differences in fluorescence, likely resulting from minor differences in staining/light exposure during the protocol, etc.
My question is: is it better to adjust for these differences during imaging (ie. change exposure and LED intensity to fit a consistent baseline fluorescence) or during analysis (ie. changing the threshold of fluorescence that is counted for analysis or the baseline amount of positive signal that is required to count the cell as positive)?
I am using a BD pathway imager. When the spinning disk gets in the light path I get very blurry images( see attachment). Does anybody know how to fix this issue. First image is with regular fluorescence image. The second is with confocal at the same focal plane.
I want to ask you a question. Do you know what kind of passivation substrate can resist DNA adhesion? I tried 5K PEG and 2K PEG modification, and found that the DNA modified with Cy3b or Alexa 488 Atto 647N will be adsorbed on the substrate through weak interaction or non-specific, and it is still firmly adsorbed on the substrate when washed with water. . Is there any other way to make DNA not tightly adsorbed under the conditions of normal optical fluorescence imaging?
I want to quantify the fluorescence image of the cells by using image j software but I am confused between the mean gray value and integrated density value and its appropriate use.
I want to use fluorescence microscopy to study fibroblasts grown on electrospun mats. Although such experiments are widely described in the literature, I faced a problem. We usually immerse the sample into a mounting medium (80% glycerol), place it between a glass slide and a coverslip, and seal using nail polish. This procedure works perfectly for cells on ordinary substrates, but the electrospun nonwovens are not wetted perfectly, and so they give rise to bubbles, cavities, and the overall distribution of the mounting medium is far from homogeneous. This problem is typical for various types of mats, made of PLA, PCL, and other polymers.
Are there any tricks to overcome this? Maybe I should change the mounting medium? Or add some sort of treatment to the mats?
I have cells expressing mvenus and mcherry and I need an intrument to quantify the fluorescence of the cells (not a flow cytometer). Can anyone please tell me if there is a fluorescence plate reader at the uni-Düsseldorf?
I have a question about quantitative fluorescent microscopy. Maybe is an obvious one, but I would like to be sure about that. I am using fluorescent images to realize quantitative measurements. I stained mitochondria in my experiments; now I am using this imagess to measure the correlation with the red channel (mitochondria) and the green one (cytoplasm), with Cell Profiler. My question is: Will I modify the intensity of the pixels within the channels by improving the contrast or/and the brightness of the images previous to analysis? In other words, could this kind of image processing altering my data?
I am a microscopy image analysis noobie.It'd be great to have a discussion on this question.We have multi channel fluorescence images of dendrites and other brain structures obtained before and after an experiment on mouse.In the images obtained after the experiment,in some channels,fluorescence from 2 different viruses are present.Is there a way to separate the two fluorescence?
When you use polystyrene plates that are not 'image compatible' for fluorescent imaging, is there a significant image size distortion? I know auto-fluorescence is a cause for concern, but are there additional drawbacks to using them?
I want to conjugate protein having C-terminus with a dye having amine derivative through carboxyl-amine reaction. Does somebody has experience of doing this reaction? OR through EDC reaction chemistry? Thanking you in advance.
I want to equip the microscope to be able to make ratiometric measurements of fluorescence in living cells. I have searched several articles, but the information is ambiguous, some papers mention a shutter ..
I stained FFPE tissue with two different colors for two different antigens. I can deconvolute the original picture in FIJI (ImageJ), assign pseudofluorescent colors to the two deconvoluted channels, and merge them back together again to make one pseudofluorescent picture with two different colors.
However, I cannot find out how to assign a third color for the spots in which the two stainings are colocalized. I tried to use the coloc2 extension, but this produces only quantifications of colocalized area and no images to the best of my knowledge. Can anybody help me or maybe advise an alternative program for this purpose?
I want to perfom life cell fluorescence imaging over several days with rather low sampling rates (1 fig/h). To facilitate tracking and signal quantification, i would like to reduce cell movement/displacement to a minimum.
Whats the best way to reach this without impairing viability, and, especially, without affecting fluorescence imaging, as I am dealing with low signals anyway?
I thought of coating with ornitin/collagin/fibronectin (...), but would be happy on any expirience in advance on those or others.
Anyone heard of biotin/streptavidin immobilization of cells on glass?
Btw, dealing with U-2 OS (osteosarcoma) cells, so specific hints are also more than welcome.
Thanks for your input!
Hello Good people
When I stained my adherent non-transfected cells with Hoechst 33342 staining it showed blue fluorescence but dull staining happened with GFP transfected cell
I used 2ug per molar
30 min incubation at RT
300ul per well in 12 wells plate
So, what's your suggestion for better procedure to be able to see cell segmentation more clearly!
What's the benefits from PBS washing as recommended by some protocols at the beginning or the end!
Can anyone give me some pointers on how to carry out an SDS-PAGE in-gel fluorescence assay with a uvGFP tagged membrane protein?
From various protocols I've come across, I understand that I can run the protein on a normal gel with my usual SDS sample buffer as GFP is stable enough to still fluorese, but I should not boil (which I would not do with a membrane protein anyway) or heat the samples in any way. I assume that I should therefore just leave them at RT for an hour or so? Some protocols have also said that it essential to run the gel at 4 degrees to avoid degradation, is this actually important?
Just on a side note, uvGFP is typically excited at 395 nm, and apparently the UV tray we have with our Biorad EZ gel-doc does give a UV source that covers 395 mm. Do you think I would be able to image the gel using that to excite the uvGFP or would I need to find a dedicated fluorescence imager? I will be using Biorad Stain Free gels with the plan being to image the fluorescence and total protein on the gel, before Western blotting for the proteins Strep tag. If I could image the gel on the same gel-doc by just switching between the UV and stain free trays then that would be helpful.
I want to measure mitochondrial calcium influx using fluorescent probe (Rhod-2 AM) in primary hepatocytes. I was wondering which is the better way to measure it e.g. fluorescence imaging or flow cytometry.
I see most of the articles used fluorescence imaging. Could someone suggest/share the protocol for flow cytometry, if possible.
I need to isolate a tiny fraction of macrophages that could successfully kill the internalized bacteria (for example, M. tuberculosis). I think the best way is FACS, but I need fluorescent dye(s) that deferentially stain these two populations, while keeping their RNA content intact so that I can perform RNA-Seq.
I found that the signals for my dye are weak in tissue sample, what is the maximum limit to which I can increase the detector gain and laser power percent for my dye?
We would like to scan protein/peptide slides. Mainly, these are microscopy-sized slides with proteins dotted onto them, and then we visualize binding of another protein to these dots, using fluorescent antibodies.
The problem is that we only have the basic model of the Typhoon Trio available, which doesn't come with a slide holder. This slide holder is needed so that your mounted slides do not touch the surface of the scanner, which would cause smearing of the dots.
Do you have suggestions how we could mount and scan the slides without smearing? Any DIY solutions out there?
Thanks in advance.
I would like to know if anyone has tried microfluidic chip experiments - particularly fluorescence imaging on an upright microscope. What are the parameters that we need to keep in mind to choose an objective as I need to work with high magnification (60X).Or if anyone has tried placing and imaging a microfluidic chip in an upside down manner on an upright microscope using some sort of adaptor so to still use a high numerical aperture objective?
I try to get confocal images of p-tau in human brain.
As shown in the image below, p-tau S396 antibody that I used is able to stain neurofibrillary tangles (Red signal) in AD patients brain.
(Blue is DAPI signal)
but I also see weak signals of dotting pattern in soma.
The brain of normal person has the same dotting-like signal too but weaker.
Is the dotting-like signal in soma is false (non specific signal)?
How is normal tau (not pathological form) stained in normal and AD patients brain with p-tau antibody?
I hardly find fluorescent images of p-tau staining in human brain.
Is it because it is better to stain p-tau by DAB staining?
Looking forward to your answer.
I currently perform primary cultures of cardiomyocytes of foetals rats (almost born) and I would like to validate that there are only cardiomyocytes and for that to make a specific and different marking between the cardiac cells. Mainly specific to cardiomyocytes
Thank you in advance if you have markers to recommend me and for your help.
I have seen some workers used Matlab in combination with Image J for that purpose, others used scripting languages as well, Is this means that ImageJ can not do all the necessary tasks alone with all its free plugins.
Hi, I want to perfuse phospholipid (PS/PC : 30/70) into a capillary and after that take some florescent images. After adding PS and PC into the glass vial, I allow the chloroform to evaporate. Then I add detergent into the vial and inject it into the dialysis cassette. I am wondering when I should add fluorescent dye to the sample? If I add fluorescent dye after the dialysis, can it be conjugated with the phospholipid?
The cells are EndoC-BH4. They are grown on matrigel, but they do not seem to be submerged into the gel. I tried to stain them with antibodies, phalloidin, and DAPI after paraformaldehyde fixation and permeabilization. The outcome was very poor. Has anybody a good experience with any cells grown on matrigel? Any advises?
Can any one provide with a recommended DAPI staining protocol for dental pulp stem cells already fixed and dried for SEM imaging?.The cells are fixed on dental chips 4*4mm.Want to use these samples for fluorescence imaging instead of SEM imaging now.
I am doing an invasion assay that uses calcein AM but the cells I want to assay contain the GFP gene and since they emit at approximately the same wavelength, I am not sure if it is feasible to use calcein or not. Does anyone have any idea how to make this work or do i need to purchase a different viability dye for the assay?
Currently I am working on a project where I inject GFP-expressed ovarian cancer cells into zebrafish embryos. I injected cells at 2dpf, and took images at 2dpi and 4 dpi. And I am tracking tumor progression by taking green fluorescence images.
However, I found a really weird thing today when I was imaging by uninjected control: clearly there are spherical regions that both emits green and red along axis of fish, but I don't understand what is causing this problem... because I am tracking green dots to verify my hypothesis and this is interfering my results...
Any thoughts on why this is happening? I am currently using EK strain and the images were taken at 4dpi.
Hi. I made a mycoplasma test with HOECHTS 33342 on MCF-7 cells and MDA-MB-321. I've used glacial acetic acid and methanol (1:3) as a fixation solution. Also I've incubated my cells with 0.5ug/mL (working solution) of Hoechst. Do you think it's overexposed ?
Name of assay, fluroscence molecule should be used for knowing the number of amine molecules immobilized.
I have a set of cells double stained in this manner: first the cells has been live-stained to visualize the protein of interest at the membrane (named: SURFACE). Then, cells have been fixed and permeabilized. A second staining was then performed for the same protein of interest (named: TOTAL).
The two fluorophores have different colors (green and red). So far I've analyzed them by normalizing the SURFACE mean intensity to the TOTAL mean intensity (SURFACE/TOTAL), but not fully happy with this way of analysis and data representation...
Does anyone knows a better way to correlate the two signals?
(PS: the main problem is that the two signals have different intensity. Red is generally more faint than green, and even if it should mark the TOTAL protein the ratio between the two is always around 1 for control...)
I needed a mitochondrial marker for immunofluorescence co-localizazion analyses. I decided to use the Thermo Fisher Scientific MitoTracker (cat. no. M7510). I stained HUVECs with 100 nM - 500 nM MitoTracker for 20 min - 40 min, and obtained nice images. As specified by the supplier, the oxidized form of MitoTracker is sequestered in the mitochondria (https://www.thermofisher.com/order/catalog/product/M7510), thus I was expecting to see a very specific signal only within mitochondria. As You can see, all my cells showed a network of interconnected mitochondria (i.e., the expected signal). However, many of them exhibited also a clear intranuclear positivity.
Here I attach some representative original b/w micrographs and false-color merged pics.
Has anyone an explanation for the nuclear positivity? Is it possible that the signal I got from within the nuclei derived from clusters of mitochondria just above of below the nuclear envelope? Would an observation by confocal microscopy clarify that?
Thanks a lot in advance
We are working on the preparation of transition metal dichalcagonide (TMD) samples with 60-90µm sized monolayer trinangles of WS2, MoS2, WSe2 or MoSe2. The goal is to cover a large area of the substrate with monolayer TMDs to perform further experiments. We are looking for a cheap and straight-forward method to get a first quick estimate of the sample quality right after production in our CVD set-up. I know that the PL quantum yield is much higher for monolayer TMDs than for multilayers. Therefore, I have the idea to put them onto an optical microscope with up to 1000x magnification and look at the luminescence. My first approach was to do the optical excitation with a laser (laser pointer, laser diode, HeNe). However, I realized that a lot microscope nowadays offer the option of LED irradiation. Is green LED irradiation sufficient to measure PL of TMDs?
Is it feasible to differentiate CD45.2 cells from CD45.1 cells in chimeric mice marrow using immunofluorescence? I need to add another 3-4 colors in order to tell different cell location and are they CD45.2 or CD45.1. Thanks in advance for any suggestion.
I need to compare fluorescence values between ROIs from different images, taken with a confocal microscope. I'm using ImageJ for this purpose.
When the images were taken, the voltage/offset settings from the instrument were different for the photos I need to compare (I'm not the operator of the microscope), so the contrast seems to be really different.
Can I make a correction or something likewise to solve this problem?
I'm looking for alternatives to Alexa 555 as a fluorescent dye to calibrate my focal volume for Fluorescence Correlation Spectroscopy (FCS) measurements. I was wondering if one can point me to a reference or paper which measured the diffusion coefficient of any particular FCS-suitable dye in this range of wavelength (excitable by a 561 nm laser line).
I fix cells with 4% PFA/sucrose for 15 min, and I block with 5% normal serum from secondary species. I recently started getting this issue, and I was able to successfully fix and stain one time without the issue.
There is some substance that is lying in between the areas with cells, which you can see from comparing the phase contrast to the fluorescent images. But, the Hoescht and green staining seems to indicate that there is some substance that is fluorescing on the cells.
Has anyone seen this happen before? I rinse with DI water before mounting the cover slips and it doesn't look like PBS crystals, so I don't think that's an issue.
I don't know why there's something between the spaces with cells that isn't fluorescing at all, but I have a few general suspicions:
1) Serum is sticking. I'm using really old serum (circa 2009) that has been aliquoted and stored at -20.
2) Cells are somehow drying out, although I aspirate and add solution in a sequential manner.
3) I use a vacuum to aspirate during washes and such, and possibly it's not strong enough to remove all the liquid.
4) I recently switched from VECTASHIELD non-hardening mounting medium to the hardening one.
My pictures are in jpg format, I need to know if it is possible to quantify fluorescence of S100B and GFAP in cells. And wich is the best way to do this, using mean gray value or integrated density? In both cases I do not have clear how the measure is, wich is the units or how can I know this? I attach a sample image for a marker, Thanks!!
I'm attempting to fluorescence image tumour microsctrutures, in particular:
However I've found most stains become unbound or degrade during ethanol dehydration (which is required before I can embed my samples in resin). Any suggestions for robust stains?
Thanks in advance,
I am running fluorescence assay for the first time, the excitation and emission wavelength of my compound consecutively are 300-410 nm, while the plates in my lab (white clear bottom plates) enable reading down 340 nm only. My question is could I use those plates for my experiment while the emission wavelength is out of the plate's reading range? I mean is it supposed to use a plate with wavelength reading's range including excitation and emission or emission only?
I am using CLARIOstar® device to run this experiment. I tried my best to look online to find plates which allow reading up to 410 or more but I could not find any. According to the original assay article, they used acryl disposable cuvettes, which is not practical for my assay as I have many dilutions for many samples and controls and I do not have a device can be used with cuvettes for fluorescence assay.
I am working on fluorescence image and want to see the intensity distribution of an image object from the same. I am using MatLab to measure it. Please help me in this regard.
I am trying to track a bacterial DNA binding protein during its cell cycle. I have phase contrast image of the cell and the protein tagged with GFP. I want to track my protein inside cell in time-lapse experiment. Is there any FIJI/ImageJ plugin that can work with both phase contrast and fluorescent images together? Thanks in advance
It doesn't matter the color of the fluorophore or the localization (nuclear, cytoplasm). I just need a good one, because the ones I tested so far didn't give great result. Any suggestions from people with experience in that. Thank you very much for helping.
I have to analyze muscle fiber stainings and dapi labelled myonuclei.
1. fiber typing to distinguish type I/type II fibers
2. assign myonuclei to fibers
3. count fiber areas
I came across Fiji and ImageJ and would like to ask if anyone did muscle fiber typing with Fiji/ImageJ before and if they could provide some help. I would like to automatize this as I would have to spend weeks on analyzing my images. Help/Hints or scripts would be highly appreciated.
I'm currently trying to do some calcium imaging using micro-manager. However, I couldn't find any plugin to monitor fluorescence intensity of ROIs over time online.
Does anybody know someone that could help me with that?
Thanks a lot,
I used an anti-ANT antibody fluorescently labelled with Alexa Fluor red and tried to show co-localization with mitochondria using mitotracker red but it did not work. Why does it not work and what is a better approach?