Confocal Fluorescence Microscopy - Science method

Hope this article helps.

I agree with Randall Scott Gieni, 300µM tissue section will not only be difficult to permeabilize but your antibody penetration will be difficult.

Good luck,

Hi, i think you can use a concentration of 2ug/mL.

Dear Fatemeh,

Maybe you look through this protocol:

Reagents

Equipment

Procedure

Timing

Troubleshooting

Steps 1-4 It is important to handle the cardiomyocytes as gentle as possible while they are viable. The optimum temperature should be maintained and low-speed centrifugation in a temperature controlled centrifuge should be carried. The pipette tips should have a wide bore (by cutting the tip of the pipette tips) so that the cardiomyocytes are not strained while transferring.

Anticipated Results

With applying the above-described protocol, it should be possible to get images of isolated adult mouse cardiomyocytes which has been immunostained using confocal microscopy. The shape and morphology of the cardiomyocytes is retained even after fixation.

Figure 1 Confocal images of fixed cardiomyocytes with pseudo-colors for each staining.

Figure 1a Isolated mouse ventricular cardiac myocyte stained with MitoTracker Deep Red 633 for staining mitochondria shown as blue, Alexa Fluor 568 phalloidin for staining F-actin shown as red and SYTO 11 Green-Fluorescent Nucleic Acid stain for staining the nucleus shown as green. The images are deconvolved using Fluoview software from a series of image stacks obtained using Olympus Fluoview confocal microscope FV300.

Figure 1b Isolated cardiac myocyte stained as above shown as X-Y view. The orthogonal projection on the left of the image is the cross-section passing through the myocyte along the Y-Z view. The orthogonal projection on the bottom of the image is the cross-section passing through the myocyte along the X-Z view (images obtained using Leica TCS SPE confocal microscope).

Supplementary Figure 1a-d Isolated mouse cardiac myocyte pseudo-colored showing different staining in different panels and the fourth image on the bottom right is a merged image of all three panels (images obtained using Leica TCS SPE confocal microscope).

Supplementary Movie 1 The above described image stacks were used to construct a 3D image to visualize the distribution of cytoskeletal proteins and organelles within the cell.

References

Sathya Srinivasan, Experimental Imaging Centre, University of Calgary

Correspondence to: Sathya Srinivasan (sathya_sr70@hotmail.com)

Source: Protocol Exchange (2011) doi:10.1038/protex.2011.235. Originally published online 11 May 2011.

Best regards,

Alexandr

You can set the min and max, for example:

setMinAndMax(2, 160);

The waitForUser(""); command will introduce a pause in the script until you click ok. You can also add a message as an argument e.g., waitForUser("manually adjust the brightness and contrast");

I work mainly with mammalian cells and have no clue about the plant cells but I feel this could help you with your DAPI problem. DAPI is not great at staining a live nucleus. DAPI most often cannot enter cell membrane/wall and staining requires fixation and permeabilization. I suggest you try Hoechst 33342 which has similar excitation and emission wavelength so you don't have to change your protocol.

Please note that this works for mammalian cells and may or may not work for plant cells but something to think about.

Dear Alexandr,

Thank you for the answer. Unfortunately, I already know these works really well!

I can't use neural crest markers in IF as most of them are either cytoplasmic or downregulated before the enteric neurons become terminally differentiated.

Sox10 works quite well up to a certain point, but it's absent in mature neurons: its expression is maintained in enteric glia only.

Propidium iodide cannot be used as a viability dye in fixed cells. Propidium iodide is usually used to measure DNA content in fixed cells, but that requires methanol fixation and RNase treatment. And even then, I wouldnt rely on the sub-g1 population as a reliable for viability assays.

If you want to know the viability of fixed cells, look into amine reactive fixable dyes such as life technologies live/dead dyes, biolegend's zombie dyes, or the ghost dyes. You must first stain your cells with one of these amine reactive dyes and then you can fix the cells without losing your dead cell staining.

These dyes tend to have sharper excitation/emission spectra as well as numerous color options. You should be able to stain for both viability and phalloidin, just avoid fluorophores with spectral overlap.

It will depends of your FRET design. If a fluorochrome such as DAPI is not emitting or absorbing the light wavelength needed in your FRET system you should not have any problem. However, DAPI is a fluorochrome with a very broad spectrum of absorption and emission so it is maybe not be recommended. You could use phalloidin coated with an Alexa fluorochrome that is not having interference with your FRET system.

You can check the fluorochrome spectra of individual dyes and their overlap with other (if any) in:

I Just checked your macro and it looks okay to me. Maybe you should try and include in the last step

run("Flatten");

I sometimes experience errors with merged images in TIFF format and this seems to help

Debaprasad Koner if you are seeing fluorescence with mitoRed then you are on the right track. Mitotracker red is membrane potential specific so what you see are mitos with reserved MP, and the stain is retained post fixation. However if you want to do a mito count then mitotracker green is the best since it is not mp dependent. Some mitos will lose MP based on treatment, mito fitness, lysis etc. MitoGreen also retains well post fixation.

Hello,

You may coat your coverslips with Poly-L-Lysin for example and let your cells to sediment and attach on the glass.

You can also click on > Analyze tab > Click Set Scale and reset the scale (this is to convert your pixels to microns where you should have a scale bar of known size).

Sathya Srinivasan

Hi Huijing, the goal of confocal is to reduce the intensity from out-of-focus planes, but the cutoff is never a sharp one, so strong signals from planes not too far from the focus can be observed in the image. Make sure that your pinhole is not too large. If larger than 1AU, you will have increased contribution from out of focus. Other factor to consider is that DAPI signals are often very strong, compared to your FP, so you may see some faint out-of-focus signal from DAPI but not from the FP.

Dear Cy,

First, be sure that your cells are not contaminated by mycoplasma because the cytoplasm is not stained by HOECHST. Second, when you observe your stained cells by a conventional fluorescence microscopy, the light dispersion causes the nucleus seems as integrated round blue units. Whereas by using confocal fluorescence microscopy, the unfavorable light dispersion will be eliminated, hence you'll observe the stained nucleus with more details. Since the HOECHST is intercalated into the DNA, and regarding the fact that chromosomes don't occupy all the space of nucleoplasm, I think that your microscopic image is absolutely perfect.

My pleasure, good luck with your work!

Hi Saulo,

Thank you for the response. I was concerning if the GFP fluorescence can be damage after the fixative method. Do you have any ideia about it.

Thanks

If you are using Imaris to open these images (which you hashtagged in your question so I am assuming), then after opening the images, imaris gives you the option to click on the image using the cursor and reposition it the way you want it.

Uncheck the "frame" option to get rid of the frame. Take a snap and open it using imageJ, and in there you could re construct the xy axis (make sure you know the pixel size before you uncheck the 'frame' option, and draw the xy axis accordingly) hope this helps.

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.

Brian Sanz

I was actually going to ask you if you had double checked your filter/Cy3 combination, but since you were already using Cy3 successfully I figured you were good to go. You might throw some of that SA-Cy3 dilution between a coverslip/slide and just make sure you can see it with your 514 set up.

I wish I was being more helpful!

Primary vs secondary antibodies and direct vs indirect staining

Primary antibodies provide specific recognition of the target antigen. Secondary antibodies bind to primary antibodies based on the species they were raised in. Primary antibodies that have been raised in rabbit or mouse are amenable to detection with anti-rabbit or anti-mouse secondaries, respectively.

The pairing of a primary antibody and secondary antibody (specific for the host species of the primary) that is conjugated to a fluorophore is known as indirect staining. The use of primary antibodies conjugated to fluorophores, without secondary antibodies, is known as direct staining. Direct staining methods save time and allow multiplexing using antibodies raised in the same host species. Indirect staining traditionally offers higher sensitivity thanks to signal amplification that occurs when multiple secondary antibodies bind to a single primary antibody.

Other variations on these techniques are available, such as using biotinylated primary antibodies with avidin/streptavidin-conjugated fluorochrome. Fluorophore-conjugated F(ab’)2 secondary antibody fragments (lacking the Fc domain) can help reduce background. In mIHC, horseradish peroxidase (HRP)-conjugated secondary antibodies are employed to catalyze deposition of tyramide-fluorophore conjugates.

Brian,

I would not recommend to use MitoSox with fixed tissue!

1. MitoSox is a membrane permeable weak acid. In live cells it distributes into the mitochondria because , as long as the mitos maintain their membrane potential, the mitochondrial matrix is the most alkaline place in the cell. Once MitoSox arrives in the matrix it gives off a proton. Now electrically charged it is no longer membrane permeable which effectively traps it inside the mitos. Thus, for proper localization it depends on the presence of membrane potentials which are missing in fixed tissue.

2. MitoSox becomes fluorescent after reaction with superoxide. The stain is meant to report superoxide as it is produced by the mitochondria, typically by activity of the electron transport chain (normal or pathologic). Fixed mitochondria are not metabolically active any more and therefore should not be able to produce superoxide any longer. Superoxide is diffusible, reactive (ROS - duh!) and therefore has a very short half-life. Thus, the superoxide that has been present at time of fixation will be long gone at the time you stain your fixed tissue slices.

Anything you find with MitoSox in fixed tissue will probably be caused by non-specific effects, such as autoxidation, and will be more confusing than useful.

I recommend to follow Alexandr's hints and concentrate on immunodetection of permanent ROS-damage to the tissue (protein, nucleic acids).

Good luck with your project!

Bernhard

For confocal imaging, if you have two probes, red and green then emmission light of green will not excite red. It is designed in a way that you will see only what you want to see. If you want to see red along with green then you have to program the scope in a way that you excite both red and green separately to see them.

If you want to do FRET, your microscope should be equipped with a different protocol for FRET.

Hi Fran,

A mild heat-induced antigen retrieval on PFA fixed frozen brain sections has been performed for some IHC assays. At 70-80 C for 30 min or at 40 C overnight in a water bath. Unlike FFPE sections, tissue detachment from the slide might be an issue with HIER on the frozen section Tried several adhesive slides for HIER, Truebond 380 slides found better tissue attachemt or use the floating section method. Good luck

Thank you all for your help :) I will read the articles that you have shared.

Other aspect to think about is the primary ab. What is the epitope recognized by this ab? Some ab are good for western blot but not for IF because of the epitope.

Hello, I'm reviving this old post since I'm dealing with a similar situation. I have a recombinant protein with 10xHis tag on the N' terminal and a Myc tag on the C' terminal. The protein is molecular weight is ~42.4 kDa. I would like to use the protein for analyzing antigen-specific memory B cells. I have in mind staining the protein with a fluorophore-conjugated anti-Myc or anti-His, and then using the labeled protein to stain the B cells population, followed by Flow cytometry analysis. Does anybody have an experience with such a procedure aimed at B cells? I am aware of the alternative of using the Molecular probes kits, but the first methods seems cheaper and more straightforward. Thank you!

There are few assays which utilize this concept for diagnosis like DHR assays. where the intensity of conversion of substrate is taken as diagnosis.

Except from this, if you talk about surface markers, many diseases are diagnosed on basis of expression of certain markers tagged with fluorescence molecule.

Sorry for the slow answer! I'm using smaller spheroids (max 500 µm) which are easier to clear and image. Maybe some clarity protocol would be suitable in your case. I would also recommend you to check the working distance of your objective if you want to image the whole spheroid. A long distance objective might be needed to image deep inside the spheroid core.

Hi there,

The answer above is all right and is more of a BatchProcessing. You have also another option which is to assign a shortcut to your Macro so you just have to press a key to perform the function. In the example attached here is "F9". Btw, it will be much faster if you decide a value for each one of the channels for B&C adjustment so you can write it in the macro, this will prevent you for adjusting it in each channel each time. If you want to use different values for different sets of images, just rewrite the macro. The macro here automatically does all what you were doing, but image by image, by pressing F9. It will save the final image in the same folder where your original image was.

To use it:

Download the *.ijm file attached here.

Go to Plugins>Macro>Install...

Then open the image and press F9 and it will be done.

It also creates a white scale bar of 10*1 um at x=2um y=2um of your image. If you want to change those dimensions or the position, just modify the values 10 and 1 in the run("Specify..." line.

One warning, this is assuming that your C1=Red, C2=Green and C3=Blue. If it is not the case you have to modify the run("Merge Channels..." line. c1, c2 and c3 are always red, green, blue. If you have more channels, you will have to add them following the same reasoning.

Hope it helps,

Cheers

macro "adjust BC [F9]"

{

title = getTitle();

filename_clean = File.nameWithoutExtension();

path_to_dir = File.directory;

path_to_save = path_to_dir+filename_clean;

run("Z Project...", "projection=[Max Intensity]");

close(title);

run("Split Channels");

selectWindow("C1-MAX_"+title);

run("Brightness/Contrast...");

waitForUser("Apply optimal Brightness and contrast settings, then press OK");

selectWindow("C2-MAX_"+title);

run("Brightness/Contrast...");

waitForUser("Apply optimal Brightness and contrast settings, then press OK");

selectWindow("C3-MAX_"+title);

run("Brightness/Contrast...");

waitForUser("Apply optimal Brightness and contrast settings, then press OK");

setForegroundColor(255, 255, 255);

run("Fill");

saveAs("tiff", path_to_save+".tiff")

run("Close All");

}

Might try a non-iterative method like Wiener if the PSF is known.

I Agree With Sayantan Mitra

I agree with Sönke Weinert . Probably the easiest is trypan blue. This is a common technique often employed with phagocytosis assays, and a technique we use with our Vybrant Phagocytosis assay kit (link below). Since the FITC is on the surface, the trypan blue works through absorbance quenching to quench the FITC signal. Since trypan blue is not cell permeant and not taken up via phagocytosis, the phagocytoses bacteria then becomes unquenched and fluoresces green upon entering the cells.

So, all you'd need to do is put a subset of the labeled bacteria into a solution of trypan blue, compared to a control with no trypan blue, and see if the signal is quenched. If it isn't quenched, then the dye is internalized in the bacteria. If it is quenched, then the dye is on the outside of the bacteria surface.

Be aware, though, that if the bacterial cell wall and/or plasma membrane are compromised, the trypan blue will also enter the cell and quench internal signal.

Trying to determine the localization of signal via high-resolution imaging is reliant upon having a very high-performance microscope or super-resolution system. Most standard widefield or even confocal microscopes are not reliably capable of determining it with enough precision:

Vybrant kit link:

As suggested by the others, ImageJ/ Fiji is a very nice tool to analyse microscopy images. If you're new to bioimage analysis and would like to get a better understanding, I can really recommend you the following channel:

Concerning comparing you samples: it it not be the best idea to compare fluorescent intensities! Samples bleach due to storage, antibody performance varies and different laser intensities/ illumination times can bias your results. Try to find a more reliable way, like quantifying cell numbers or marker-positive area.

This paper could be interesting for you: DOI:10.1038/s41598-017-16797-1, however, the there used ImageJ plugin is currently broken. But it can give you some ideas about possible acquisition and evaluation steps and perhaps the issues get fixed soon.

If you are having troubles with certain analysis steps, you can find help here: https://forum.image.sc/

Happy imaging!

If you have a single channel grayscale image there's probably nothing you can do to get the original color channels. Such image contains only the information about intensity averaged across all three channels per each pixel.

If you change the format of your grayscale image and treat it as an RGB, splitting will give you the same image for red, green and blue, all identical to the original grayscale image. This is because only a mix of these 3 colors in equal and specific intensities results in the same corresponding intensity of white = certain shade of gray in every pixel ("grayscale" image).

If you had an RGB image made by merging of original red and green images, each pixel would still contain information about the intensities of red, green and blue channels separately. You would get the original channels by splitting the RGB image with one difference: If the original data were e.g. 12- or 16-bit, splitting an RGB image will provide only 8-bit result (ImageJ doesn't work with higher quality RGB in default, there might be some plug-ins), which means that you are losing information and quality. You could discard the blue channel which would be all black = zero intensity values in your case. I just tried all of this with one of my images and it functions the way described. Note that this method can only be used if the original image was made of red, green and blue channels or their subset, nothing else!

Anyway, this should be the last emergency solution to be used only when

a) you work with an image from someone else (watch for copyright) and you don't have an access to the original data or

b) you've lost your own original data.

Of course, the best practice is to keep your original data before any processing (and possibly some intermediate processing steps) ideally with a back-up and use those.

A Ferreol mentions, DeconvolutionLab2 is mainly a learning tool. However if you are processing only 2D images it could be good enough for your needs. If you are processing many, large 3D images the speed could be an issue. It is possible to use GPU deconvolution in ImageJ, though right now that requires an advanced set up to get imagej to use the fast GPU math and FFT libraries (if interested ask about it on the ImageJ forum https://forum.image.sc/ ). It looks like you are having issues dealing with a multiple channel image. In this case each channel would have a slightly different PSF. To process them properly you need to split the image into separate channels, generate a PSF for each channel, then merge them back after deconvolution. In ImageJ you may need a specialized script to do the entire workflow.

DAPI stains RNA as well, unless you pre-treat the cells with RNAse. Maybe try Hoechst (33458?) because it does not bind to RNA, just to DNA (it binds to the "rungs" of the DNA ladder, which RNA doesn't have).

Use fluorescent conjugated secondary antibodies ex. Alexa fluor conjugated antibodies

Are you looking for 3D information like volume and surface area? If so, you need to analyze them as a volume with 3D Objects Counter. If you are looking for 2D information, then you don't need the 3D Objects Counter plugin and you can do standard 2D measurements. If the projection isn't too filled with structures (which can happen in large z stacks filled with dense structures), then I prefer to measure in the sum intensity projection rather than the maximum intensity projection. A max projection is only selectively displaying some brightest pixels from the volume and skews the measurements. The Sum projection shows the integrated value of all pixels, so especially if you are quantifying intensity this is important. or, if the information you need is contained within individual planes then you can measure a representative plane or planes.

It seems that it would depend on the species of cyanobacteria.

Hi Fan,

Sectioning and staining of lymph node is not different from other tissues.

But you know that thickness of lymph node compare to other tissue. Therefore, you have to make very thin slice of tissue. If you already inject dye labeled nanoparticles then you can just use nuclear stain and visualize under confocal microscope.

Many thanks for your help.

The best results can be achieved with high-pressure freezing, freeze substitution and flat embedding.

Good luck!

Hi. If you are planning to use primary cells, it is better to establish your own stable cell lines. I have 2 recommendations. 1) Choose actin-GFP, not GFP-actin because attaching GFP before actin seems to affect cell's behavior (https://www.jstage.jst.go.jp/article/csf/42/2/42_17016/_html/-char/en)

2) If you are trying to establish a stable cell line, try lentiviruses (better pre-packaged and ready-to-use, such as from here https://www.systembio.com/products/imaging-and-reporter-vectors/cyto-tracers/pct-actin-gfp-cmv/), or go with transposable elements (piggyBac plasmids).

This answer is copied from earlier discussion on researchgate under similar question that might help you

By Daniel Lee Adams   If your going after a surface, extracellular, protein then there is no need to permeabilize. If the protein is intracellular or transmembrane then you need to permeabilize. Saponin, tween and triton are all good options use 0.1%-1%. Methanol or ethanol are also options if all else fails. However all proteins and cells are different, start with saponin or tween-20, then move on to the triton, then as a last resort ethanol, then methanol. This is also the order of stringency, so the saponin/tween will do the least damage to the proteins/lipids, while methanol does the most damage.

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!

Have you tried BTS?. It works in skeletal muscle inhibiting the ATPasa activity of Myosin heavy chain.

If you want to stain live cells, formalin fixation is not possible. After 2 days in formalin everything will be dead.

I could imagine that it depends on the cellular position of the antigen you want to stain: Staining cell surface proteins will be much more likely than proteins sitting in the mitochondrial membrane for example as you need to get access to the epitope.

Good luck, it sounds like an interesting but difficult idea!

Some news :

It works really well for me with Hoechst under 405 excitation, with very few toxicity on one hour visualization. I used low laser power (can't remember how but less than 1%).

Note that I work on confocal zeiss LSM 780.

Thanks all for your answer, find attached a picture of my staining:

[url=http://www.noelshack.com/2018-46-3-1542190510-hoechst.png][img]http://image.noelshack.com/minis/2018/46/3/1542190510-hoechst.png[/img][/url]

(Hela Cells)

Have a good day

Hello, here the info page, you can find much more than you looking for. https://www.olympus-lifescience.com/en/microscope-resource/primer/techniques/confocal/fluorophoresintro/

Gustavo:

I would suggest using the Photon Counting Histogram (PCH) ; see the seminal papers below.

PCH and FCS are determined on the same time series of events. To extract the distribution of the duration of the fluctuations we use a math based on calculation of the correlation function, or FCS. To extract the distribution of the amplitude of the fluctuations, we use a math based on the PCH distribution.

FCS provides the diffusion coefficient D and the number of particles <N> in the observation volume; PCH provides the brightness ε ( the number of photons emitted in the unit time per molecule) and the number of particles <N> in the observation volume. The brightness may change in your system depending upon the number of fluorophores you have.

1. The Photon Counting Histogram in Fluorescence Fluctuations Spectroscopy"; Chen Y1, Müller JD, So PT, Gratton E. ; Biophys J. 1999 Jul;77(1):553-67.

2. Characterization of Brightness and Stoichiometry of Bright Particles by Flow-fluorescence Fluctuation Spectroscopy. Johnson, J.1., Chen, Y., Mueller, J.D. ; Biophys J., 2010, 99(9), 3084-92.

Hi. Can you please expalin what you need help with? Are you trying to quantify the percentage of your cells which was affected with the drugs Or?

With more info I might be able to help.

The word you are looking for is "dynamic range". The dynamic range in your images seems to be too small to see the weak signal (peroxisome) without saturating your strong signal (plastid). If I understood you correctly.

There are several ways you can try to improve on this. First, what bit-depth are you using, usually you can set the readout from the PMT to 8 or 12 bit, 16 bit on some systems. While this not increase the dynamic range as such, it subdivides it into more steps (256 for 8 bit, 4096 steps for 12 bit). Higher bit will make the files larger and might affect acquisition speed, but might give you more details on your weak structures.

Another option is HDRi (high dynamic range imaging), which you might know from your phone. In principle this can also be applied to microscopy images. A series of exposures is taken with different settings and then combined into a single image by software. Some microscope software has this function integrated but it can be done externally (e.g. using imageJ). However, be aware that this is a non-linear operation and the resulting image will not be quantitative anymore.

Hi Mona and Muneeb. Thank you so much for your suggestion.

I was wondering if the filaments with microfluids chambers are axon, I could differentiates axon. I took the thick filaments which are at proximity of soma (cell body) as dendrites. So I am confused in the selection appropriate axons. In the image I have attached, soma in not under focus as I scanned the image with prim focus on neurites (axon and dendrites). I have already taken and scanned images for only cell body(soma). That's why I need to distinguish axon in the image I have attached. Your suggestion would be highly appreciable. Thank you once again.

Hi,

It's depend on the nature of the sample (If was fixed or not)

I recommend the use of hoechst 33342 or Sybr GreenI.

Thanks Timothy for the inputs.

I'll see what works best for my samples as I need to be heating my samples at 80C for 30 min.

Regards

Hi. I have used Prolong Gold when imaging samples using STED and it has worked very well. There is minimal sample distortion and it is readily available. Make sure to follow the manufacturer's advice with regards to the curating period (>24h) and to use high quality coverslips.

Thanks for all the inputs!

I used ImageJ as suggested.

If anyone sees this question later, looking for help, here is my Excel sheet.

If you just put a drop of antifade, and then gently put the coverslip upside down to allow to sink by the viscous antifade. It will be OK.

Regards

When you used FITC to conjugate the graphene oxide, did you notice any change in the color or still its native color ??

for my knowldge that FITC is  high reactive with minor amounts of free amino groups. so to ensure the deficiency in conjugation , you must measure the content of the dye in your produced conjugate and should be following the recommended ration to get you the preferable fluorescence.

Dear Dr.Subir Das,

I have studied Efficiency of the second-harmonic generation with the zeroth-order bessel-profile beams in uniaxial crystalsand and I have also learned how to create the zero order Bessel beam. To create the zero order Bessel beam we can use annular slit as in the work of Prof.D. MCGLOIN * and Prof. K. DHOLAKIA.

However, the most convenient method is probably to use an axicon.

No one is interested in measuring the surface area in both attached and suspended cells. Due to tremendous change of cell niche in both conditions, stress, cytoskeletal system, metabolic events, the resulting signaling system received within the cell will be totally different. Therefore, it is pointless to discuss, so, no papers. Just imagine monomeric actin molecules and dimeric tubulin molecules either polymerize or depolymerize in a brink of time within the cells. This cell movement accompanies with different metabolic activities. The membranes are not a fixed boundary, and endomembane system is always flowing from the plasma membranes, cis-, medial-, and trans-Golgi membranes, and ER membranes to the nuclear membranes and ever-changing. Full appreciation of these basic biological systems can only tell that there is no surface area change in both states of cells, but changes occur in cytoplasm. Just like a completely air-leaked balloon and maximally inflated balloon!!! Still doubt?

You may be interested in the followings;

Cell 16 (4), 909-918 (1979)

Biotechnol Prog 9 (4), 362-365 (1993)

20

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.

You can use calcein AM. 

Stable: DNA is integrated into host genome, cannot be lost. Both daughter cells receive a full genome copy, complete with transgene.

Transient: DNA is on a plasmid, and can be lost (if a cell divides and only one daughter cell gets plasmid copies, the other daughter lineage has now lost the plasmid forever).

You are using transient transfection, so your DNA will eventually be lost by your cells.

You don't specify whether your plasmid contains a eukaryotic resistance cassette or not: this is a common method to ensure plasmid retention. If, for instance, your plasmid confers neomycin resistance, then your cells can be grown in neomycin and any that lose the plasmid will die.

if you don't have a resistance cassette, then...yes, your DNA will eventually be lost by your cells.

It shouldn't happen that fast, though, and the CAG promotor is (in my hands) generally very high expression and pretty robust, so even if your plasmid is lost, your cells should remain detectably green for several divisions (the green signal will simply halve with each division).

I am unclear as to why you would WANT to isolate a clonal cell population if you suspect they might have kicked out your plasmid anyway, however.