Science method

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.
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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!
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This was clearly not the edge effect in cell culture as it happened during staining procedure. It looks like the cell in the middle of the well has been washed away. To understand when the issue happened, you shall take a look at the cell under the microscope after every single steps of the staining, then you will know which step could be wrong.
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Hi everyone.
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.
Stefano Falone
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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?
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Hi there and sorry for the late answer,
To add to the answer of Christopher B O'Connell
I usually find this method the most accurate and unbiased to remove background. The theoretical explanation abovementioned by Christopher B O'Connell and the references provided are perfect.
I find Rolling-Ball advantageous because:
I) Minimizes user-intervention (potential sources of error and bias)
II) Since it's local, different values are subtracted to every pixel. Usually this is the best way to correct biomedical images unless you can assure a perfectly even illumination, which is often not the case (not due to the imaging system, but due to the sample)
Another option that some people use to remove background is to average several background ROIs (usually cell free regions, tissue free regions or un-labelled tissue) and perform an arithmetical subtraction of this value.
You can do this over the image in ImageJ by:
Process>Math>Subtract...
and input your value
Or, alternatively, after listing the values, remove this value to all your ROIs.
Again, I'd never advice this method since the user intervention could introduce errors (not perfect sampling of all the cell/tissue free regions; (un)intentional bias towards different experimental sets,...)
As a plus, in ImageJ you can find a plethora of different filters and de-noising tools (Under Process>Noise and Process>Filters, respectively). Although these aren't strictly speaking background removal tools, they can help to improve your images if used correctly. All of them are well explained here:
Hope this helps,
J
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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.
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You can use Real-time potency assay for CAR T cell killing of adherent cancer cells.
Please refer to the link below for more details about the assay.
The eSight assay couples the simplicity, analytical sensitivity, and objectivity of real-time impedance monitoring with highly specific readout of live cell imaging to characterize CAR T cell-killing efficacy.
Hope this helps.
Best.
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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.
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John Griffin. Well, you are certainly free to believe that "most people have no idea what they're doing".
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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.
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Hi James,
The opacity shouldn't matter since you are illuminating as well as observing from the coverslip side, so neither excitation nor fluorescence need to propagate through the disk. We image biofilms on steel plates in a similar way. Is there water between the coverslip and the PVC disk?
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Greetings everyone,
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,
Florian J.
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Hello everyone,
Sorry for late response, it's been some busy two weeks with other more pressing matters on hand. I'll do my best to answer the questions that came up tomorrow when I have access to my data again. (This is just a quick PSA so you dont think I didn't bother to check here again after getting an answer.)
Have a nice sunday and until then
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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.
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If it is not a confocal microscope, this is due to the fact that you are seeing unfocused light emitted from outside the focal plane.
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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)?
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To anyone reading this in the future,
I consulted with a microscopy specialist in our department and he recommended not changing the settings on the imaging step, but rather setting a baseline difference in fluorescence by comparing unstained control groups between replicates, and adjusting the data based on the difference measured.
Thanks for all your responses! I appreciate the assistance.
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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.
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In addition to the previous answer also try to increase the speed of the spinning disk if you can find an option to do that.
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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?
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I fully agree with Alexander Dräbenstedt that only mathematical postprocessing can give the solution to your problem. You can use a multichannel fast Fourier transform (FFT) preprocessing as well.
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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?
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It is possible to add CaF2
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I have a liquid that glows when UV light is shined. However, when I use my phone camera I can't see the glow, I just see a violet hue. What is the best way of capturing the fluorescence-glow from our samples? DO I need to use a filter for my camera?
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it depends on the sensitivity of the camera with respect to the different wavelengths. I expect that a mobile phone camera is a standard Si detector, hence the sensitivity at low wavelengths is very poor. For sure the use of some filters can help you to obtain better resuts, but first of all you need to know the spectrum of the emitted fluorescence (in principle you can know it if you know the dyes in your system).
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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.
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Hi there,
This has been discussed elsewhere
But (From ImageJ manual):
1) Mean gray value (MGV) Average gray value within the selection. This is the sum of the gray values of all the pixels in the selection divided by the number of pixels. Reported in calibrated units (e.g., optical density) if Analyze▷Calibrate…↓ was used to calibrate the image. [...]
2) Integrated density (ID) The sum of the values of the pixels in the image or selection. This is equivalent to the product of Area and Mean Gray Value.
So
3) Mean Gray Value*Area = Integrated density
Indeed, there is no clear-cut answer to your question, it basically depends on what you are measuring and which biological significance have your results. Both are correct, but it depends on what you want to find.
To use a parallelism in physicochemical terms: To me, this has always been something like the difference between amount of a substance (Integrated Density in images) and concentration of a substance (Mean Gray Value in images) considering volumes (that would be areas in 2D images). You can get the same amount of any substance with different concentrations, or the same concentration with different amounts (playing around with the volumes)... right?
If you assume that the areas/sizes of the particles you are comparing have similar distributions, then you can use MGV.
If your treatment (or the difference between the groups you are comparing) involve changes in the size distribution of the particles you are analyzing (cells, ROIs,...) then it might be convenient to use ID or at least consider that some differences could be hidden in MGV and arise when using ID.
Moreover, if you are using a confocal system, then you will also have to take into account that you are working in a 3D space. So the so called area, actually comes from a maximal projection of a 3D space and then, the proper way to do this is considering the 3D space, not just the area (voxel vs pixel). However, even in this case, you'll have to consider that z resolution is not as good as xy resolution in your system.
If the later is the case, you may want to take a look at this:
Hope this helps,
Cheers,
J
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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?
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The microscope is not magical ,without a good sample you won't a good sample you won't have images that are suitable for image analysis but a good design of your experiment could save you alot of time and money .Here are just highlight that have to be taken into consideration when you are preparing your sample for floresent imaging
(1)choice of coverslip
In general ,you should always use #1.5 coverslip (0.17mm thickness )as most microscope objectives are designated to work optimally with these .this is true for fixed samples .for live cell imaging ,lbidi as a very good choice of dishes and chambers .
(2)Choice of your florophores .
(3)Fixation and permeabilization e.g formaldehyde /paraformaldehyde methanol ,All these are what preserves cell morphology depending on specially .
Mounting and recording medium
Mounting medium helps preserves your sample and raises the refractive index to give good performance with oil performance.mountants often have scavengers which soak up free radicals and reduce photo bleaching these sometimes reduce initial briggteness of your samples.A glcerol PBS mixture is suitable for most fixed immunofluoresence and GEP -labelled samples .
Although they are also commercially products available e.g flouro amount -G ,Aqua/poly -mount ,pro long ,vest a shield immunity.
(4)Clearing methods
(6)Recording medium ,l will go in details in this.
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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?
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Du findest mich unter www.foranthrop.com
Beste Grüße Wolfgang
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Hello,
Does anybody have idea about auto-fluorescence of polymers and networks? Is it normal that polymers emit fluorescence?
Regards
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Hello,
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?
Thank you
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You can also try procedure of histogram equalization that allow to see variations within mitochondria that appeared uniform in the original image and to check the amount of non-mitochondrial staining. This is not useful for quantative imaging.
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Dear experts,
Liquid carbon dots (CDs ) were synthesized with greenish color under UV-light. However, after mixing with PVA polymer, the color turned to blue with enhanced quantum yield (QY). what is the possible interaction made this enhancement? Is it related to hydrogen bonds of PVA or other reasons. Thanks in advance.
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Look carefully at your data. since You list quantum yield, I assume the color is fluorescence. Was the original fluorescence green or a combination of blue and yellow - but now the yellow is gone? If it is a shift in the color from green to blue (higher energy, greater difference between the excited and ground states), either the ground state is stabilized or the excited state destabilized.
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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?
Thanks!
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Yes, of course. Plastic bottom is not transparent enough, which leads to disordered reflection lights and distorted images. It is more obvious when the images are captured at high magnification(60x or 100x). You'd better use an glass bottom plates.
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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.
Kind Regards
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If there is one possible labeling site, add at least 10-fold molar excess of labeling reagent over protein (or peptide) for maleimides. After about 2 hours reaction time, remove the excess dye by gel filtration chromatography (for a protein) or reverse phase chromatography (for a peptide).
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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 ..
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First you need a good excitation source with same power for YFP and BFP or CFP.
LED are good but not perfect. The best is xenon lamp. Xenon has the same power for all excitations (in visible).
1) Expensive solution :
Dual bands emission filter with dual bands dichroïc mirror, with filters wheel in front of the source (xenon). You need a very high quality filter.
It's slower than with LED's because LED do not need filters. You can add a DualView system and acquire the image pair in the same time.
2) cheaper solution :
2 classics dichroïcs cubes. Good but slower.
Others solutions dear colleagues??
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Hi Everyone,
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?
Cheers,
Jasper
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Make each channel a binary mask, take the overlap of the two binary masks, use that itself to mask the original non-binary combined image. You should be left with a masked out area of the original image with full dynamic range, but only in the areas where they are colocalized. Then pseudocolor to whatever you'd like.
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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!
Christian
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Hi Christian, you might try adjusting the confluence of your cells. Although you are using cancer cells, if they don't 'like' to crawl over each other then once they bump into each other they might just stay in a very limited area and allow for tracking the same single cells over long periods of time.
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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!
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Recently Im using your suggested reagents and concentrations but does it make difference if im using for example BSA 3 %
Thanks :)
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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.
Thanks,
Samuel.
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Hi, in case anyone wonders how this evntually panned out, I can tell you that I was able to get in-gel GFP fluoresence to work eventually. Essentially you just need to make samples up as normal in loading buffer, but then add protease inhibitors and just leave them at room temp for 20 to 30 mins with occasional light vortexing. Pre-chill the SDS-PAGE running buffer, but other than that run the gel as normal (I used 200V for 40 mins rather than our usual 300V for 17 mins though to limit gel heating). I was able to occasionally see a faint signal with Bio-Rad Geldoc EZ and the blue plate, but eventually found we had an Amersham Typhoon imager elsewhere in the building, which was able to show a robust signal with its 488 nm laser line.
One word of warning, initially I thought I could see expression of my protein on the in-gel fluorescence, but mass-spec protein ID could only find the GFPuv fusion tag. It transpired that the GFPuv was dimerising under the high concentration it was at, so I would recommend putting a monomising mutation (such as A206K for GFP) in the GFP tag to prevent this.
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Dear all,
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.
Thanks
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Dear Gary, Thank you for the response. Interestingly, I had already visited your lab website while searching of the calcium flux measurement in the mitochondria. It has a nice calcium profile using Rhod-2.
Your suggestions are really helpful. I am looking forward to do my experiment.
Best
Pavitra
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Hi,
What kind of automated microscopes for fluorescence imaging are you using and why (pros / cons) ?
Thanks
Francois
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We are using a Vectra 3 from (former) PerkinElmer, in between the whole department was bought by Akoya Biosciences. Has nice acquisition and analysis software included, however, I would suggest to buy the Polaris (successor of Vectra). Have no information about their other systems. No contact with the Akoya staff so far, but apparently, they took over the support team from PerkinElmer which was great. Multiplex stainings are cumbersome to establish but can be quite powerful.
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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.
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Hello,
can you have access to fluorescent bacteria? In combination with a marker for activated macrophages you could sort 3 populations:
- non activated macrophages non fluorescent
- fluorescent activated macrophages with fluorescent non degraded bacteria
- fluorescent activated macrophages without fluorescent bacteria (degraded)
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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?
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Hi Chem,
unfortunately, percentages do not mean much, since the actual power on the sample strongly depends on the light source you are using, the objective you are employing, and the other settings of the microscope, so you have to experiment a bit with your settings.
As a general guideline, photodetector gain should be kept quite high when you are using your confocal microscope in live mode to find the focal plane and image area you want to image. Just turn it up until you start seeing visible noise in dark areas of the image, this way you know you will not miss any fluorescent object just because the gain is not high enough. Once you find the are of your sample you are interested in, turn down the gain to try to limit noise as much as possible, in order to get a "good" image. Ideally you should lower the gain until dark areas are completely black, and slow down the scan speed and/or increas the excitation power until the bright areas are almost at the maximum possible signal (but be careful not to saturate the image).
As for the excitation power, it really depends on whether your sample is fixed or fresh and living. If it is live, the lower the power, the better, as exposure to high intensities of light can interfere with the biological processes you are trying to observe. If the sample is fixed, however, the limiting factor is photobleaching, meaning the more you acquire images, the more you "burn" the fluorescent molecules, and the less bright the next image will be. I would suggest, when you are just looking at your sample to find the area you want ot image, to keep the power as low as possible, as long as you see something, and only turn it up once you start acquiring the images you want to save.If you are collecting a 3d stack, keep the power lower, as photobleaching happens even in the planes above and below the one you are imaging. Finally, if you can spare the time,slowing down the acquisition speed or averaging more images will in general always lead to better results than simply turning the power up, so try to be patient and only increase the power as a last resort.
Good luck!
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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.
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Did you come up with a solution to this? I am facing same problem
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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?
Thanks
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High NA usually means short working distance. This is often problematic for fluidic devices that are intubated on one side and mounted on something rigid, like a glass slide or coverslip across the entirety of the other side.
We have used custom fabricated acrylic holders for this purpose in the past. They allow you to have a path for the tubing to exit out one or both sides while allowing your objective to come as close as necessary to the glass which is now on the top side. The setup is essentially like a really thick slide (10-12mm). Keep in mind that some objectives have working distances of less than a typical slide (1mm) in which case you'll have to mount your fluidic on a cover glass instead.
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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.
Thanks!
Jisu
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One thing to consider is auto fluorescence from lipofuscin. The distribution of perinuclear granules in what appear to be neurons (based on size) is consistent with lipofuscin. If the signal shows up mostly in the green and red channels it is probably autofluorescence. Older cases and Alzheimer's disease cases are loaded with lipofuscin. There a few things you can do to remove this type of autofluorescence. You can stain with Sudan Black or TrueView, which masks the signal of lipofuscin. You can also do some form of linear unmixing if your microscope has a spectral detector. There are some tau antibodies that do stain nuclei, but the perinuclear distribution of this material is most likely autofluorescence from Lipofuscin.
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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.
Tiphaine
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For Fibroblasts:
For Cardiomyocytes:
Cardiac Troponin T
Cardiac Troponin I
Sarcomeric Alpha Actinin
Tropomyosin
Best
Harsha
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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.
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ImageJ is a Java-based image processing program with a lot of useful plug-ins, where the one or the other might fulfill your requirements. However, you did not mention what exactly are your individual tasks in these images. I have experience with Matlab and Python. If you have a licence for Matlab you will have less stress with libriaries and dependencies. If you have no licence, Python is reasonable too and you can install and work wherever you want. If you need algorithms using artificial intelligence (object detection, automatic (semantic) segmentation, in Python there is probably more code and repositories available. I would only recommend ImageJ or other standalone (bio) applications (e.g. Icy - Bioimage analysis) if you never want a program a line of code.
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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?
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I assume you are using the detergent removal protocol to make liposomes? It is not the best nor the easiest method if you do not have protein in your sample. Even if you have no extrusion possibility you could use the ethanol injection method and skip the detergent addition, dialysis, and unavoidable detergent contamination, and have your liposomes in seconds.
I would prefer using a fluorescently-labelled lipid derivative (e.g. Bodipy-PC) that you could add in the very beginning, even if you dialyse it afterwards. But you can of course add a relatively water soluble probe after the dialysis if you wish.
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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?
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What are you using to permeabilize? I'd try 0.5% Triton/PBS for 15 mins, wash and stain incubating for a longer period of time and/or at a higher concentration of DAPI/phalloidin. I'd expect at least one of these will show an improvement from your first round of staining and you can optimize from there.
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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.
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I'd just rehydrate with any suitable solvent (MeOH, EtOH, etc) and PBS and once it's in PBS DAPI stain in the usual manner. I'd suggest 1/10,000-20,000 for 10-15 mins followed by 3 x 5 minute washes with as much PBS as can fit into the well/container/whatever your staining it in.
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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?
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Hi! I have the same problem, but i do not measure viability with Calcein-AM but Pgp-transporter function. So the dye switch is not so simple for me (Calcein-Violet and Blue is not Pgp substrate). Any idea? Thank you!
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Hi all,
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.
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These are developing pigment cells; the stripe along the dorsal surface is quite characteristic. They autofluoresce in both the green and red channels, and you can see this in the image– most dots have both colors.
To remove this, you must inhibit pigment development from 1 dpf on (we use PTU solution) or use an albino strain of fish. And one must always take comparable images of negative controls (same stage, no injections) using the same illumination.
Finally, I would check on your injected cells much earlier. At 2 dpf the zebrafish already has a working innate immune system, which means thousands of macrophages and neutrophils that can kill foreign cells. Unless you are inhibiting the innate immune system or using a immunocompromised strain of fish there may be no injected cells alive by the time you are trying to see them (48 or 96 hours later). So you need to confirm that the injected cells are viable and for how long. I would inject and then image much earlier-- such as 1 hour, 2 hours and 4 hours, for a start.
Good luck with your experiment.
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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 ?
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Hi Lucas,
The question of if something is overexposed seems to really be relative to what you are looking for. Since you are looking for mycoplasma, I don't think that this is overexposed, your nuclei are nice a bright, but the puncta of Hochts positive areas surrounding at the periphery of the cells are still visible. I am not exactly familiar with these cells, but it looks that there is a modest level of mycoplasma contamination (more so in the MCF7 than the MDA-MBs) so I would consider treating your cells with BM cyclin 1 and 2, or, even better, throw them away and start over with fresh aliquots of early passage cells. That being said, Matthias is right, PCR is the most definitive test for mycoplasma contamination. I hope that this helps.
Best,
Andrew
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In a comparative manner I wanted to know the sensitivity of fluorescence imaging, MRI, CT and PET.
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Name of assay, fluroscence molecule should be used for knowing the number of amine molecules immobilized.
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You could use the commercially available reactive fluorescent dyes which are routinely being used for attaching fluorophores to proteins (extrinsic fluorophores). These dyes react specifically with -NH2 or -SH of -COOH groups present on the amino acid side chains. From their fluorescence, you could quantitate the number of free NH2 groups. I hope the -NH2 groups are free.
There could be some complications due to quenching, dimer formation and excimer formation. Choice of the dye is going to be critical to avoid these complications.
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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...)
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Im curious as to why you don't stain the cells post fixation with both sets of markers. A live cell will indeed react differently to fixed cells. If you did a serial staining procedure you can still image total protein and surface:
1) block the cells and stain with your surface marker and it's corresponding secondary. (no permeabilization)
2) Permeablization step and stain other primary and secondary.
What antibodies are you using? Different antibodies will have different levels of effectiveness in staining (as I am sure you are aware) and could cause a discrepancy in your results. If they are the same with a different fluorophore attached, you may be able to correlate better. There are lots of kits that allow you to conjugate antibodies. You could then compare intensities of staining for surface and total with each color to normalize and make a comparison. What is the colocalization signal like? Could you quantify the intensity of the yellow?
Good luck!
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Hi everyone.
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
Stefano Falone
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Just to throw an extra monkey wrench into the conversation here...
In grad school we developed a "mitochondrial" sensor that was using triphenylphosphonium to localize but it also showed very clear nucleolar staining. I confirmed it was nucleolar by confocal co-localization with nucleolin-GFP and fibrillarin-GFP. This is all unpublished (and probably never will be), but we originally thought it might be that the moiety our sensor was developed for was also in the nucleus.
I think there must be some charge distribution specific to the nucleolus. It would make sense, considering how much nascent RNA Is synthesized there that there would need to be a lot of cation for stabilization - perhaps there is a unique charge distribution there.
Here is another example of published work with TPP (and mitotracker green) that went to the nucleolus (figure 2): http://www.mdpi.com/1424-8247/10/4/91
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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?
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Following
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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.
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Thanks! Will give it a try...
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Hi!
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?
Thank you!
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Hi Matias,
comparing intensities in microscopy samples is a non trivial problem. Acquiring images with the same settings is a must and even then there may be variations due, for example, to laser power instability or sample thickness and preparation. I think many underestimate the errors that can be introduced. In my experience, the best way to do this is to find an internal reference to compare to You could use another protein that you know is present at the same level in all your samples or a dye or if, for instance you are measuring a membrane protein you can measure a cytoplasmic to membrane intensity ratio and then compare that ratio between samples. Of course it depends on the question you are asking and on what your sample is. However, if the difference in acquisition settings is considerable I would, personally, not trust the data for other than an indication...
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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).
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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.
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In my esperience is the situation 2). some times I observed the same phenomenon when the cells were drying S......
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assay is based on conjugation antibody to measure T cells proliferation
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Thank you so much dear Elisa
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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!!
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Hello,
  1. Select the cell of interest using any of the drawing/selection tools (i.e. rectangle, circle, polygon or freeform) (TIFF images)
  2. From the Analyze menu select “set measurements”. Make sure you have AREA, INTEGRATED DENSITY and MEAN GRAY VALUE selected (the rest can be ignored).
  3. Now select “Measure” from the analyze menu or hit cmd+m (apple). You should now see a popup box with a stack of values for that first cell.
  4. Now go and select a region next to your cell that has no fluroence, this will be your background. NB: the size is not important. If you want to be super accurate here take 3+ selections from around the cell.
  5. Repeat this step for the other cells in the field of view that you want to measure.
  6. Once you have finished, select all the data in the Results window, and copy (cmd+c) and paste (cmd+v) into a new excel worksheet (or similar program)
  7. Use this formula to calculate the corrected total cell fluorescence (CTCF).
  8. NB: You can use excel to perform this calculation for you.CTCF = Integrated Density – (Area of selected cell  X Mean fluorescence of background readings)
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I'm attempting to fluorescence image tumour microsctrutures, in particular:
blood vessels
cell membrane
cytoplasm/cytosol
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,
Natalie
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You could flush the vasuclar system with a biotin solution that'll stick to the blood vessels when fixed. Biotin can be easily visualised with staining with streptavidin coupled to a flourescent protien.
You can also inject biotin into individual cells.
You could also flush the system with CM-Dil. It is a lipophilic dye, that will stain cell membranes. It is a very bright and dye, that is fixable and will survive the harshes conditions, even a week in 8% SDS and clearing with glycerol or ethanol. We used it in CLARITY, see attachement.
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Hi:
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.
Thank you
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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.
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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
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The single molecule tracking technique helps in your case. Please find the following links that might help you. The link 1 should be more close to your case. Goodluck
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There is a consensus about this? I couldn't find any.
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JTV-519, dantrolene and ryanodine at high concentrations are options but i have no idea which would be the best choice
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AX
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It seems that you have another emitter, if what you show are colocalization images. What about an absortion of the second emitter on the QD ?
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Chromosome counting using fluorescent microscope....
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If you want a good imaging of chromosomes with both conventional fluorescence microscopes, semi-confocal technologies and/or confocal microscopes (even super-resolution and nanoscopy microscopes) then forget about DAPI mounting medium, specially Vectas..... this will cause an high background, diffused two-photons emission that will coock your samples and inefficiency in your overall imaging. I'd like to suggest you to use Hoechst 33342 or picogreen for DNA visualization (compatibly with other fluorocromes you need in your preps), stain, wash, rinse and mount with a DAPI-free mounting medium. Optical index is something unknown by most but really important for imaging. Your mounting medium OI should match those of your coverslip and immersol (immersion oil) if used. Glycerol is really good (90% glycerol with clean PBS is fine). Antifading and antiquencing is extremely important to me so bear in mind that. Other commercial mounting medium are extremely well made in that sense. Also Mowiol powder could be used to make an excellent mounting medium but as well as glycerol you must add an antifading at least. Always clean coverslips and never autoclave them, they're not pirex glass and they'll be coocked.
feel free to ask any other question.
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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.
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You can use calcein AM. 
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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.
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Cytoinformatics.LLC company has extensive experience in muscle fiber imaging analysis. It includes fiber area analysis, fiber-type-specific CSA and myonuclei counting and much more. Based on deep learning and advanced computer vision techniques, the analysis is 100-160 faster than the current procedure. 
Contact me at       chong.liu@cytoinformatics.com      if you are interested. 
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Hello,
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,
Fernanda
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Dear Fernanda,
In addition to Zhilin's answer, you will find all required sources of information by following this related issue on ResearchGate: https://www.researchgate.net/post/How_to_measure_mean_fluorescent_Intensity_MFI_from_confocal_images
Best Regards
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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?
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If you can micro-inject the antibody into the cell, then you can use it along with the vital stain of mitochondria. But as John Hildyard pointed out, you need to use two different colored dyes to do the colocalization study.
An alternative approach would be to stain fixed cells with your anti-ANT antibody and a second labeled antibody (with a different color) that also recognizes a mitochondrial antigen.
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I wonder if someone used the imageJ to analyze the puncta-like fluorescence intensity using Image J software? I tried to quantify the intensity using regular method, but the S.D. appear too high, the regular CTCF (corrected total cell fluorescence) method might not be proper for the puncta-like staining. Who knows how to quantify it? the picture i attached is the one i hope to quantify. Thank you.
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Dear Mou. The columns you use to set a calibration, for example: if what is black in the image represents 0 mg of calcium (just an e.g.), and the strongest red is 25 mg of calcium. In this case, you can make a calibration (in this case using only 2 points, the zero and 25... ). So, as black is represented in ImageJ (8 bits images) by zero, and white (red in the case you choose it in the lookup table) is 255.. so you can make
in the left column:
0
255
in the right column:
0
25
and choose the straight line for the function. 
After calibrated, you can select an ROI (region of interest) by choosing the forms (rectangle, circle, etc.) available, and then go to Analyze>measure (CTRL+M) and you will have a statistics of the region you selected.  You can also go to SET Measurements to select what do you want to measure...
Hope it is what you are looking for...
All the best
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I would like to be able to compare fluorescence measurements taken at different times in different samples directly with one another.
From previous experience I know that microscope settings need to vary between samples in order to optimise the image being taken. I use live tissue and variations in dye loading, tissue thickness, etc. prevent keeping the microscope settings (detector gain, amplifier offset, amplifier gain) the same between measurements of different samples. Obviously settings for different measurements made within the same sample are kept the same, and laser power and exposure time are also kept constant for different samples.
As such, is there a way to normalise fluorescence measurements? Ideally I'd like to know If I can normalise my fluorescence measurement of interest (e.g. TMRM) to some cellular autofluorescent species (something that shouldn't vary in my experimental conditions of course). Something essentially akin to the loading control used in Western blotting. I'd measure both my probe of interest and the 'loading control' under the same microscope settings (just varying the excitation and emission wavelengths as necessary). Is this at all possible? Or does having to change excitation and emission wavelengths make this impossible.
Alternatively, could I normalise measurements of the fluorophore of interest from my region of interest (I currently take measurements in my time-course experiments by selecting the nephrons in kidney tissue) to the background fluorescence of the same fluorophore? And if so what counts as the background? Interstitial tissue? The perfusate fluid that surrounds the tissue on the stage?
Any other suggestions? Some kind of physical standard like a fluorescence ruler, or fluorescent beads?
I do not want to manipulate the data with ImageJ or some other software as I think this will somewhat invalidate it.
Many thanks for your time!
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I doubt you could have a source of autofluorescence that you could just assume to be stable and unchanging. If you have not yet done the experiments, I would suggest using fluorescent beads with wavelengths that are close but not overlapping.
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I have thousands of  images that fall into 7 classes, I wanted to try using Relion 2.0 but I have no background in coding, does anyone know if fluorescence image data in TIFF format can be used by or converted to a suitable form for use by Relion 2.0?
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In this publication (http://science.sciencemag.org/content/341/6146/655/tab-figures-data), they have used IMAGIC single particle pipeline to process the Super-Resolution Microscopy data. 
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I have several slides of cells stained for transcription factors and cytoskeletal proteins. The staining worked quite well and appears very specific to targets. Now I'd like to have some semi-quantitative analysis of the slides. What is a typical method of analyzing stainings? For example, can one us an arbitrary flourophore intensity to bin cells into either a 'high' or 'low' expression level, and count the number of each type per field of view? Do people try to produce an average intensity for the FOV? What is common here? Please link to papers if possible, thank you!
Update: Attached is an example field of view. These are cancer cells which I have probed for vimentin (red) and e-cadherin (white). All cells in every experimental group express both proteins to a degree and I am attempting to  quantify potentially subtle expression differences.
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I agree with Alex. Cell profiler will enable you to count cells, assess colocalization of your antibody staining pattern, and measure intensity values across your images very easily. The best part is it will batch process hundreds of images while you go for coffee! And it's free!
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Co is the initial concentration of the dye, while Cn is the concentration of the dye after dilution. Intensity of the images are found using ImageJ as well as MATLAB, there is hardly any difference in the grey values obtained from both. But the curve shows, otherwise, ie. increase in concentration with dilution, which is quite puzzling. Please help, Thank you
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In simpler way it can be explained: As the concentration of solution increases there is increase in factor called internal quenching.
This internal quenching occurs due variety of processes like excited state reactions, energy transfer, complex-formation, collisional quenching, etc which get increased with increase in number of moles in the solution i.e. with in concentration of solution.
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