Confocal Microscopy - Science method

Hi Oishika Chatterjee

Probably You should try the Prolong Antifade reagent from Thermo (Link Attached). in my experience I have observed that it gives better signal than the Vectashield, Fisher and Sigma's.

I have had some successs treating it like a protein gel asset. I use a small spatula get in between the seal and then slowly twist to crack the well from the slide. You may need to do that in a few spots before fully detaching the wells. This has worked most of the time, but I have had a few break on me if I use too much force. I hope that helps.

Hello,

When performing the APEX2-Gold method on animals, there are a few things to consider:

Overall, the APEX2-Gold method is a powerful tool for detecting and analyzing APEX2-tagged proteins in electron microscopy. With proper technique and high-quality reagents, you should be able to detect a strong signal in your samples.

Regards/

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

I would try confirming the expression by western blot using both myc antibody and antibody for the protein of your interest. The existence of mRNA does not mean that protein was expressed.

Myc tags are not ideal for microscopy because endogenous c-Myc could also be detected and could interfere with the analysis but it usually doesn't.

Malcolm Nobre Hello, many thanks for your answer.

If the dye is lipophilic (lipid-attracted), then how does it bind to hydrophobic structures like Golgi membrane using hydrophilic (water-attracted) groups? If the dye is lipid-attracted (lipophilic) can it have hydrophilic groups?

Kayli Rose Jablonski I suggest to read these articles:

1. https://labanimres.biomedcentral.com/counter/pdf/10.1186/s42826-020-00044-2.pdf

2. https://www.nature.com/articles/s41467-020-19827-1.pdf

3.

4.

5.

6. https://protocolsmethods.springernature.com/posts/combined-whole-mount-fluorescent-in-situ-hybridization-and-antibody-staining-in-zebrafish-embryos-and-larvae

7.

Hello Deven Gopinath

You should avoid using blocking serum from the same host species as your primary antibody that is detected by a secondary antibody against the same species (immunoglobulins in the blocking serum will compete for the secondary binding).

I agree with Yulia Panina. Washing is an important step. Washing after each application of antibody or other fluorescent probe eliminates antibodies with lower binding affinity present in the sample and thus reduces non-specific signal, or cross-reactivity. Washing for a few minutes in PBS with at least two buffer exchanges will help to eliminate unbound and loosely bound antibody from the sample.

If you are using donkey secondary, then you should use donkey serum for blocking which you have already been doing. Instead of donkey serum you may also try using using BSA (3-5%) for blocking.

Best.

Increasing the size of pinhole will definitely increase the background. I would suggest to keep the pinhole at 1 or may be increasing the pinhole size to 1.5, but in addition, I would definitely suggest to take multiple z-stacks to get a composite image with bigger signal and low background. Hope this helps. Good luck.

Thanks Both Jiasong and Ayse for your answers!

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.

Thank you Yulia Panina, I did more tests and the results are better (no tail and green ring around the red droplet), thus it seems to be matter of sample preparation.

Probably if pH is too high, the FCF dye loses its property.

Many thanks for your interest.

Best wishes,

Filippo

Thank you Alibek, I will do it and I hope it will work :)

Yes. Here is the protocol that we used for detection of proteins on the surface of Lactococcus lactis bacterial cells.

20 μl of cell culture in stationary phase (OD600=3) was added to 500 μl of Tris-buffered saline (TBS; 50 mM Tris-HCl, 150 mM NaCl, pH 7.5) and centrifuged for 5 min at 5,000 × g and 4°C. The pellet was resuspended in 500 μl of TBS, and 1 μl of FITC-conjugated human IgG antibody (Jackson ImmunoResearch, West Grove, PA) was added. After 2 h of incubation at RT with constant shaking at 550 rpm, cells were washed three times with 200 μl 0.1% TBST and resuspended in 300 μl TBS. Stained cells were fixed to a microscope slide by use of a StatSpine Cytofuge 2 centrifuge (Iris Sample Processing) for 10 min at maximum speed. An LSM 710 confocal microscope (Carl Zeiss, Oberkochen, Germany) was used for observation of prepared specimens. Alexa Fluor 488 was excited with an argon laser (488 nm), and the emission was filtered with a narrow-band 505- to 530-nm filter. All images were taken with the same settings and were analyzed using Carl Zeiss ZEN Lite 2012 software.

Hello,

once you do image treatment, you can define a background treshold on the read signal you find as background. You can also do a Z-stack to enhance your bead signal and also have bead signal in another stack that you might be missing. You can merge your stacks to have the maximum signal. If not, you can also do FACS and seek for double tagged populations ^^

Dear Omar,

do you got the answer for this question? I've meet the same situation and tried to find the explanation.

Thanks.

Hi Diego Suarez have you been using this probe at the end?

I think that you can use the Golgi staining to measure the density

Alexa Fluor 647-coupled antibody, Alexa Fluor 488, Alexa Fluor 500, and Alexa Fluor 514 may might be helpful.

Thankyou

Hi Maria,

I agree with other people that EPCAM and E-cadherin are good markers. The staining will be found at the lateral membranes of cells (E-cadherin has strong signal at adherens junctions and weak signal at lateral membrane). ZO-1 is also a good marker for tight junctions although it also stains blood vessels and mesothelium (You can distinguish them by morphology).

When you optimize your protocol, I would reccomend to try different fixation method. Most antibodies of tight junction proteins give great staining with ethanol (or methanol) fixation. Acetone treatment following ethanol fixation sometimes improves S/N ratio.

Good luck!

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

Mohamed Khashan

As has been mentioned, a simple sub-cloning should do the trick. The caveat being that you have some DNA carrying the coding sequence of mCherry.

Cloning options:

1. Restriction enzyme-based cloning

2. Restriction-free cloning - rf-cloning.org (this works great and is often cheaper, especially if you don't want to order restriction enzymes)

3. Gibson cloning

Happy cloning!

Dear Teena Bhakuni,

If you visually cannot distinguish single&full cells, even with lower magnification, then I am afraid that you need additional staining that evenly stains the membrane to quantify cell surfaces (as I have mentioned above) so the boundaries between cells are clear. Also, nuclei staining would also help to distinguish single cells. PS: General rule: one can only quantify things that are visible.

Regarding the delineation of membrane and cytoplasm compartments - same rule. You can only measure/delineate if you see it, so I am afraid that lower magnification than what you showed above, would not allow that. Also, as already mentioned you have no marker for the membrane, so the delineation will not be very accurate. You would need additional staining of the membrane on top of your protein of interest to be accurate.

Furthermore, is there a way to seed the cells at lower density? This could greatly help your analysis as maybe then you can easier distinguish single cells.

Good luck!

Mounting media usually contain anti-fading agents, so they help prevent photobleaching. If it is too thick, you could dilute it with some PBS. Of course you could also use plain PBS, if you are in a hurry and you fo not care about keeping the sample in good condition for later. Ideally you should put the medium on the one side, place the coverlip at a 45 degree angle, then slowly lower it so that the medium goes everywhere on the tissue with as few bubbles as possible.

I would say that the question doesn't stand Okolab vs. Tokai Hit. There are many solutions from other companies as well.

Recently I wrote a blog post for FocalPlane about temperature control in light microscopy. I think it could be useful for some people reading this thread.

disclosure: I work for a company that produces a temperature control device. It's called VAHEAT and it could be the device for you, if you do a lot of high resolution microscopy, need high temperature precision, need temperature cycles, not just a constant temperature, use microfluidics and more. interherence.com/vaheat/

Yes. It is matter of concern and although alternative methods are available in order to prevent oocyte damage during handling in vitro. The alternative methods have some or other type of negative impact on the oocyte. The development of an improved method/materials that can replace glycerol and prevent damage under experimental condition is required. To the best of my knowledge no such appropriate wax cushion is in the market which can be better in handling oocytes in vitro or for the assessment of maturation process in mammalian oocytes for confocal microscope.

Hi Marija,

People tend to do one of two things when determining sample size. You can either refer to a number of replicates that are „conventionally accepted” in your field for a particular experiment, or alternatively use statistical calculations to determine an ideal sample size based on certain assumptions/preliminary data. In the absence of standard protocols, you could go down this second route. For this you’ll need to have some idea of what an „average” set of results from your experiment look like. The number of replicates and fields of view per replicate will depend on:

I would recommend you have a brief read (or watch this YouTube video https://www.youtube.com/watch?v=clbg10IwzYA) about power / sample size calculations so you get the theory, and then try an online calculator such as the one found here: https://www.stat.ubc.ca/~rollin/stats/ssize/n2.html

For the first point, the number of fields of views, the main question is whether your 15 cells you currently image accurately represent the entire well. You would need multiple fields if for example you notice that within a single well there are regional differences. If Mitoxantrone intensity is fairly evenly distributed I would not worry too much. If you are concerned, I was wondering if you could extend your field of view. If you quantify average intensity in the whole cytoplasm, you could probably get away with using a lower mag objective to capture a larger field of view and higher number of cells.

Regarding the variability between experiments, if you have already performed it a few times you should have an idea what the standard deviation is and you could plug that into the sample size calculator.

You have to count manually this way, which you may want to do on a regular fluorescent scope first then use confocal to collect your publication quality images. You will quantify this by calculating the percentage of Annexin-V positive and PI-negative cells divided by the total number of cells. Flow is calculating by sorting and is not necessary. Also, something to think about, are you trying to differentiate between apoptosis or secondary necrosis?

The simplest explation is that your sample is drying out. As the buffer becomes more hypertonic RBCs undergo crenation. https://www.thoughtco.com/crenation-definition-and-example-609188

Dear C. B. , the simple and general answer to your first question should be clearly negative, the cellular uptake and cellular location of NPs are different experiments. However, they could be the same experiment if all the uptook NP´s are localized at the same organelle or cell region.

With respect to your second question, if you have two different drugs encapsulated in a polymeric delivery system to facilitate the uptake by the cell, they could be detected by confocal microscopy if these two drugs show distinctive fluorescence signals. It means that these two drugs must show auto-fluorescence in the conditions of excitation used and they must show different "colours" to be distinguished. If one or both drugs do not show fluorescence under the working conditions or they show the same or very close colours, then you must use some dye to label each drug. If your delivery system is intended for reaching some cell´s organelle and just there delivering its cargo, may be it could be possible to just follow the whole system (drug A + drug B + polymeric system) and labelling it with the addition of some fluorescent dye, well inside or attached to the polymeric shell. In this way you could see if the whole system reaches the cell and then the target organelle. To use this methodology you need to be sure that both drugs reach the target and that the polymeric shell keeps its cargo till the destination, in other case you would be following an empty- fluorescent- shell.

Hope this helps. Good luck with your research work.

In apoptotic cells, the membrane phospholipid phosphatidylserine (PS) is translocated from the inner to the outer leaflet of the plasma membrane, thereby exposing PS to the external cellular environment. Annexin V is a Ca2+ dependent phospholipid-binding protein that has a high affinity for PS. Since annexin V staining precedes the loss of membrane integrity which accompanies the later stages of cell death resulting from either apoptotic or necrotic processes, staining with annexin V-FITC is typically used in conjunction with a dye such as PI allows to identify early apoptotic cells (PI negative, annexin V-FITC positive) from dead cells (PI positive, annexin V-FITC positive). you can see this picture for easy understanding- https://www.creative-bioarray.com/support/annexin-v-apoptosis-assay.htm

If you want to distinguish late apoptotic cells from necrotic ones you have to use flow cytometry. You can also check this for reference

Dear Aedrian,

We have faced your problem in the past too - but if you have a good microscope and camera system I think that you can usefully and quantitatively examine differences in biofilm structure using some common dyes for bacterial cells and EPS.

The key questions are whether you have a fluorescent microscope and camera system. If you do, then I'd suggest looking at using live/dead stain mixes, or propidium iodide, and whichever fluorescent DNA and protein dyes you can get. If you only have a light microscope, then you need to be using standard coloured dyes, again for as many cellular components as you can. The second question is whether your camera system can capture single (black and white) or multiple colour images.

No matter what type of microscope and camera system you have, you should be able to take good images of biofilms at a resolution from 10x - 40x and perhaps even 100x (at which point bacterial cells should be visible). You can import these image files into free software such as ImageJ - you can then divide the image into squares and get the total absorbance (e.g. for live/dead stain) and use this in your statistical analysis.

You could look at the variation across the image and correlations between different stains to investigate heterogeneity. You can compare mean values for each image (or measure the absorbance for the entire image) to then compare across images (e.g.different incubation conditions, different strains, etc.).

Regards, Andrew

Thank you so much.

Hi there,

The main difference of using freeware vs built-in colocalization tools in expensive software is the price :)

Basically, these software incorporate algorithms to express colocalization and pack them into an user-friendly interface, but they're in no way superior to any other (free) software or plugin.

Additionally, you can gain insight into what these software are doing, so you will be able to know which analysis suits better your experiment by using freeware and reading carefully the guides and the manuals (Thresholds, background subtraction, ROIs, and so on...)

I have always used ICA with pretty good results (it operates in a similar manner to JACoP) and you can define ROIs in it. However there are many others. Check the attached documents and links below for practical working examples and explanations. The two attached documents are for ICA, but they provide good working examples and differences for the coefficients

Overall interesting review:

JACop Plugin original paper:

ICA Pugin original paper:

ImageJ forum about differneces in JACoP and ICA:

Why to choose one or another coefficient?

Manders' Original paper:

Guide to ICA:

Another free plugin:

And, for the second question:

No, the requirement is not for the software, but for the explanation/justification and interpretation that you give to the analysis and results. Usually, all the Plugins incorporated in ImageJ have been published if that's what you mean

Why you have chosen this analysis? Have you corrected the background? Are you using ROIs? Is this analysis powerful enough? Why you provide one coefficient and not another?

In this case, understanding what you are doing and using proper controls should be enough to validate the analysis tools that you are using for colocalization.

As positive controls, using two secondary Abs against the same 1ary Ab is usually a really good positive control (if you have time and can spend Ab's doing so). You can also perform randomization to see how accurate your coefficients are, but all these things are explained in the attached documents,

Cheers,

J

Does anyone know how to insert a scale bar on an image using evos xl core? This option is not shown to me on the screen. I’m not sure if it’s not updated or if I have to set this somewhere.

Any ideas?

If you were to image in an inverted cconfocal microscope, you would need a proper holder that can hold a 96 well plate (depending on what magnification you want to image at). However, you can only image one well at a time. Hence, I would recommend (and you would get much better quality images on) using MatTek dishes that have a glass bottom cover slip.

You can add ‘spacers’ as mounts to raise up the coverslip. Use a diamond scribe glass cutter to cut slices of coverslip. Mount these on your slide using nail polish or glycerol in parallel so your coverslip will rest on top of them with your sample in between in mounting media. Then seal round the edges with glycerol gelatine.

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!

You may also contact Thermo Fisher Scientific: https://www.thermofisher.com/sa/en/home/life-science/protein-biology/protein-expression/mammalian-protein-expression/fluorescent-protein-vectors.html

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

You may use the following sentences:

Confocal microscopy was used to take representative images of glucose uptake. These images were obtained by utilizing excitation/emission wavelengths of 488/516 mm.

There is also a nice online tool to calculate the Nyquist rate:

Hi,

Is this a spinning disk confocal you are using? If so then try increasing the speed of the disk, because the pattern seems like it probably could be forming due to the spinning disk speed being slow. You can get upgradation on the speed.

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.

Out side my experties

Hi, I was wondering how this project ended up? It seems like a good idea but I wasn't sure if the resolution/signal strength would work out.

If you are good in Python/R there are couple of scripts available. If not, try ImageJ- depends on what depth of morphometry analysis you want to do. You can also use NucleusJ plugin in Image J. Check these paper-

Good Luck !!!

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 Ahmed,

I assume you are using a confocal microscope, so you should be able to use any emission wavelength you want, as the resolution should mostly depend on the excitation beam. If that is the case, check the specific excitation spectra of the green-yellow beads you find. While the maximum of absorption will probably be at 488, beads are generally very bright, and have quite wide excitation spectra, so you should be definitely fine using them with a 440 laser, and probably even at 405.

As for the bead size, you should be fine as long as it is smaller than the diffraction limit (wavelength divided two times the NA). The very important part is for them to be very bright, not necessarily big, so in case try to slower your scan speed as much as possible for the measurement.

Also, let me introduce to a very important secret in microscopy: While for published data you should always use the proper calibrated beads from a reliable source, for internal testing you can try to dissolve the ink of Stabilo highlighter pen in water (it has to be Stabilo branded, it never worked for me with pther brands). The ink is slightly hydrophobic, and will form very bright sub-diffraction aggregates. That's a good source of very cheap "microbeads" in different colors.

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!

No as I remember I only see expression in the epidermis.

It makes sense that very high overexpression lead to aberrant localization but you still can see the normal localization (nuclear envelope in your case) which is logical.

Any way I will try to reduce the agros concentration it might reduce the expression level.

Thanks

Please check out my paper on multiphase complex coacervate systems on my profile. I detail a Confocal method for measuring fluorescent molecules inside such droplets.

Confocal microscopy can as well be used to image surfaces or internal structures by recording the scattered or reflected light intensity. Because the light intensity is much higher than with fluorescence, this alows for a much higher scanning speed. Care must be taken that the detectors can cope with these intense light powers. PMTs must probably be protected using a small pinhole setting and minimum illumination beam intensity or a filter.

José Carlos Quilles Jr You can try the following protocol. Also, if you want to use coating reagents such as Poly-L-lysine, you can coat 0.1 mg/mL of PLL (in autoclaved water) for 5 minutes by shaking. Wash this with autclaved water and dry for 2 hours. You can then add the cells there and fix them. Fixed cells can be imaged after multiple days or week (if the fluorescence is strong enough)

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");

}

Thank you all for your quick reply. I started to check them out one by one. Mr. Dirk Luetzenkirchen-Hecht than you but usnforturately Gwyddion does not have hybrid parameters command for the measurements I need (Sdr. Sds, Str, etc). I search and check many time with no results. Digital Surf is also have this lack of parameters unforturately Mr. Márcio Mafra. Thanks again for all.

It work with both of them.

You should be able to use Fiji (imageJ) for doing these. I use the plugin called QuimP on Fiji ( you can get the QuimP guide online) and it works pretty good for me to do segmentation and analysis (to be precise it gives you area, fluorescence (both cortical and cytoplasmic), circularity, elongation analysis followed by a heat map too if you want to generate one.

You can calculate the chromatin condensation parameter (CCP) simply by staining the cells with DAPI.

Thanks Khadija Elboukhari for your excellent insight.

Qiran Chris I do not have the machine available, while thanks for your answer.

Dear Tommy,

You can use deblurring and sharpening algorithms to restore the quality images / video. If necessary I can give you the codes.

Try this website:

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).

Dear Yulia , Thank you for your answer , don't u use mounting media after Dapi staining ?

Usually with a CMV driven vector transiently transfected in to a cell line you will get a huge range of fusion protein expression levels (usually beyond the dynamic range of a single image with most imaging platforms). The expression level will depend on the fusion protein half life, the time after transfection, - or more accurately nuclear access- and the number of copies of the vector introduced into cells. Cells will only start to express protein from the vector after mitosis and the levels will then tend to increase untill the cells are fixed.

Even if your endogeneous protein is cell cycle regulated in abundance then you should still get a good linear relationship at the top end of the expression levels, the regression line will just not go through zero. The non-transfected cells in your population will give you a good idea of the dynamic range of the cell cycle regulated endogenous protein levels. Incidently, after a transient transfection (say after 24hs with fugene6 ) transfected cells usually appear in pairs with similar expression levels. The distance between the pairs of cells can give you a crude estimate of the time since mitosis.

Phospho specific antibodies are more of a challenge - you could try expressing a phospho-mimetic mutant vector to see if that works with your antibody, if it is specific for a particular phosphorylation site?

We once managed to get a really good polyclonal phospho specific antibody to a serine in I kappa B alpha that on western blots was much more sensitive (at least 10x ) than the antibody to I kappa B alpha itself, which just goes to show how varied antibody affinities are.

The problem is converting to a Z-projection. This approach will superimpose cell nuclei and cell processes from multiple levels, making a pa