Fluorescence Imaging - Science method

I misunderstood your question, I thought you are using this indicator:

but actually you must be using this:

And your aim is not to measure pH, but to track the labeled proteins intracellularly. I don't think that repeated exposure of cells to excitation light according to the plan you mentioned will affect the results.

You obviously have to consider that if your protein escapes from the endolysosomal system, phrodo fluorescence will significantly decrease:

May consider using Permai fluorescence dye.

In method 1, do you have a separation step to remove unincorporated (excess) dye? If the fluorescence intensity of the dye is about the same whether or not it reacts with the protein, there will be no detection unless you remove unincorporated dye.

Method 2 provides the separation method, namely SDS-PAGE. This method will definitely work. The unincorporated dye will probably run at the front. If the dye concentration is too high, it may streak through the lane, in which case you can soak the gel in methanol-acetic acid destain, with a few changes of the destain solution, to remove it and decrease the background.

If I understand the growing condition correctly, you are going to let your cells adhere on top of collagen and then suspend the cells with matrigel? If you are sure that your cells can remain adherent on the coated plates, I guess you can use dispase to digest-off the matrigel and then only to proceed with your standard staining protocol. Otherwise, it seems that people do proceed with fixation right away with the right fixative agent.

This paper I shared belong to a senior whom I learnt organoid culture from. He basically suspended cells into 1:1 matrigel:medium mix for his organoid and then performed immunofluorescence staining straightaway by 1:1 methanol:acetone fixation. If I remember correctly, due to the solvent nature of the fixative, he did not need to worry about the matrigel. For your case on using paraformaldehyde, there is a mix experience based on the inputs on several people from this discussion:

So, now you have options whether to dissolve or not dissolve the matrigel. It seems that if you want to use paraformaldehyde, you need to pay attention to its pH and concentration.

Good luck!

May consider using Permai fluorescence dye.

For whom it may be of interest, here is a very helpful answer from Krunalkumar Shah:

There are several software options available for analyzing fluorescent imaging data apart from ImageJ. Here are a few popular ones: 1. **FIJI/ImageJ**: Although you mentioned excluding it, ImageJ's extended version FIJI is worth considering due to its wide user base and extensive plugin library specifically designed for image analysis. 2. **CellProfiler**: This is a free, open-source software designed for high-throughput image analysis. It's particularly useful for tasks like cell counting, object identification, and intensity measurements. 3. **Imaris**: Imaris is a powerful software suite for visualizing, analyzing, and interpreting 3D and 4D microscopy data. It's commonly used for tasks like colocalization analysis, tracking objects over time, and quantifying fluorescence intensity. 4. **Volocity**: Volocity is another software package designed for 3D and 4D image analysis. It offers features for colocalization analysis, object tracking, and measurement of cellular dynamics. 5. **MetaMorph**: MetaMorph is a versatile software platform that supports a wide range of microscopy applications, including fluorescence imaging. It provides tools for image analysis, object tracking, and time-lapse analysis. 6. **CellProfiler Analyst**: This is an extension of CellProfiler designed specifically for machine learning-based analysis of large image datasets. It's useful for tasks like classification, clustering, and data exploration. 7. **Huygens Software**: Huygens is known for its advanced deconvolution algorithms, making it suitable for improving image quality in fluorescence microscopy. It also offers tools for image analysis and visualization. Each software has its strengths and may be better suited to specific types of analysis or workflows. It's often helpful to try out a few options to see which one fits your needs best. Many of these software packages offer free trials or open-source versions, so you can explore them before making a decision.

May consider using Permai fluorescence dye.

May consider using Permai fluorescence dye.

May consider using Permai fluorescence dye.

May consider using Permai fluorescence dye.

The green signal outside the mCherry-expressing cells is most probably from flavins. Its intensity looks a bit higher than I would expect to see considering similar power densities for all fluorescence channels. But we do not know exactly what laser sources were used. If the green laser was an Andor HLE (2 W), then its power density at "50 ms/20%" would be more than 10-fold higher than that from an ILE 637 nm laser.

Concerning the green signal co-localized with the red one from mCherry, I am pretty unsure that it could be attributed to flavins. I'd rather suppose that you observed fluorescence of the mCherry proteins with incomplete chromophore maturation. It is a widespread phenomenon among RFPs (see, for instance, here 10.1007/s43630-021-00060-8) to show some minor green and/or blue populations. Moreover, fixation procedures can likely chemically modify the red chromophore and thus lead to appearance of green fluorescence.

Hi Masoud,

I would try with some autofluorescence quencher like True Black or Sudan Black. Applied before doing the staining should reduce the background without affecting the intensity of the target areas.

Thank you both! Labkit thus far seems to be working the best for a trainable segmentation.

The decision of where to put the mCherry fusion tag would depend on the specific protein being studied and the goals of the experiment. Generally, adding a tag to either the N- or C-terminus of a protein is a common approach for subcellular localization studies. A flexible linker can be used to connect the tag and protein, which may help to minimize any potential disruption to protein function. Ultimately, the best approach would depend on the specific protein and experimental design.

Agata Hajda Bidyut Das idyut Das I am imaging the fluorophore inside animal.

I would suppose that the fluorescence intensity mean for a region is the average pixel intensity for the region, in other words, the sum of the intensities for all the pixels divided by the number of pixels. Therefore, you do not need to divide by the area of the region because the area of the region is the same as the number of pixels in the region, so the division has already occurred. If you want to compare the total intensities of two regions, you would have to multiply their average intensities by the number of pixels or area of the regions.

Dear Monalisa Bettim ,

It is important to consider both the preservation of the cells' morphological and physiological features and the preservation of fluorescence in your cells.

Fixation in glycolaldehyde (GA) is a commonly used method to preserve the morphological features of cells in microscopy, as it crosslinks proteins and stabilizes cellular structures. However, fixation can also affect fluorescence signals and quench chlorophyll fluorescence.

In general, it is recommended to avoid fixation when studying fluorescence microscopy of chlorophyll, as it can interfere with the fluorescence signal. In cases where fixation is necessary, it is recommended to use a gentle fixative such as paraformaldehyde or methanol, which have been shown to have minimal effects on fluorescence signals.

Additionally, it may be helpful to perform control experiments to determine the effect of fixation on your particular Symbiodinium cultures, as different species and strains of Symbiodinium may respond differently to fixation.

Finally, I'd suggest to look for previous discussions on similar topics.

Dear Lorena Rincon ,

The Filkins paper you have mentioned provides a method for normalizing exposure times in fluorescent images by using the dark pixel intensity values. This method can be applied using image processing software such as ImageJ, which is a free and open-source image processing software.

Here are the general steps you can follow to apply the method to your images:

It's important to note that this method assumes that the autofluorescence is constant across the images, and that the dark pixel intensity is directly proportional to the exposure time. If this is not the case, the method will not work and may lead to inaccurate results. Additionally, the method is just one way to tackle the problem of exposure time normalization, and it's important to validate the results obtained by this method, by comparing them with other methods and by analyzing the images with different parameters.

How did you stain the slide with CYtoID for immunohistochemistry?. I did stain then cell in solution then i cytospin and fix the cells but it didnot work .

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.

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

Hello David John Coe

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.

John Griffin. Well, you are certainly free to believe that "most people have no idea what they're doing".

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?

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

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.

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.

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.

It is possible to add CaF₂

Hi there,

This has been discussed elsewhere

(https://forum.image.sc/t/mean-gray-intensity-integrated-density-and-raw-integrated-density/4983/3

https://www.researchgate.net/post/Is_integrated_density_or_mean_grey_value_better_to_quantify_fluorescence2)

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:

(https://imagejdocu.tudor.lu/plugin/stacks/3d_ij_suite/start

https://imagejdocu.tudor.lu/plugin/analysis/3d_analysis/start)

Hope this helps,

Cheers,

J

Du findest mich unter www.foranthrop.com

Beste Grüße Wolfgang

Dear all, if the field of interest is specified would be better. This is very common in the study of functional polymers. These links deal with this subject. My Regards

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.

In simple cases there is the possibility to separate the information channels by mathematical postprocessing. If for instance your measured intensity AM contains 100% information of true fluorescence distribution A and 10% of fl. dist. B, and measured channel BM contains 100% of fl. distr. B without crosstalk, then you can calculate A=AM-0.1*BM. In the general case it is a linear system of equations that can be solved.

Nonlinear effects (saturation..) and noise will limit the purity that can be achieved.

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.

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).

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??

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.

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.

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

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.

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)

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!

Did you come up with a solution to this? I am facing same problem

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.

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.

Marlon De Ita

For Fibroblasts:

Cardiac Troponin T

Cardiac Troponin I

Sarcomeric Alpha Actinin

Tropomyosin

Best

Harsha

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.

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.

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.

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!

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.

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

Hi,

Read this annu .Rev

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.

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!

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

Following

Thanks! Will give it a try...

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...

You can refer this article

In my esperience is the situation 2). some times I observed the same phenomenon when the cells were drying S......

Thank you so much dear Elisa

Hello,

see details in: https://sciencetechblog.com/2011/05/24/measuring-cell-fluorescence-using-imagej/

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.

FYI