Immunofluorescence - Immunofluorescence - Science method

There are a lot of unknowns here... but specifically 2years in PFA the brains are well fixed, and antigens are most definitely cross-linked, which will require a minimum of a quality antigen retrieval step. The other question should be addressed prior to starting; were these animals just decapitated and immersion fixed in PFA, or did the previous group perfuse them with a saline/PBS flush followed by PFA? IF they didn't do this it is probably not worth the following efforts because anything you get will be compromised by the presents of blood in the tissues. IF they did the perfusions removing all the blood, you may want to try to carefully extract the brains from the calavera and begin a cryo-protection protocol (sucrose and eventually a Glycerin/Glycol solution). This is a critical step; you can't just freeze the extracted brains without cryoprotection. After which you will be able to section for frozen IHC. At this point you still have questions... depending on your target antigens you are severely limited due to their age in PFA. In other words, there are many pitfalls with this attempt and your outcomes will vary greatly depending on a number of unknown variables. Good luck.

Detecting both mRNA and protein in the same sample using a combination of Hybridization Chain Reaction (HCR) and Immunofluorescence (IF) is a powerful technique, but it requires careful optimization to ensure that both signals are preserved and can be distinguished. Here are some recommendations for setting up your protocol:

Fixation:

Paraformaldehyde (PFA): PFA is commonly used for fixation as it preserves both cellular structure and protein antigens well. It is suitable for both HCR and IF. Fixation with PFA is typically done at 4% for 10-30 minutes at room temperature or overnight at 4°C.

Methanol (MetOH): Methanol can also be used for fixation and is particularly good for permeabilizing cells, which is important for mRNA detection. However, it can sometimes cause protein denaturation. If using MetOH, it’s often followed by a rehydration step in PBS.

Permeabilization:

For HCR, you’ll need to permeabilize the cells to allow the probe to access the mRNA. This can be done with a higher concentration of detergent like 0.5-1% Triton X-100 in PBS after fixation.

Ensure that the permeabilization step is gentle enough not to disrupt the protein epitopes.

Blocking Solution:

The concentration of blocking solution can vary, but a common starting point is 5-10% normal serum (from the species in which your secondary antibodies were raised) or 2-5% BSA in PBS or TBST.

The blocking step is crucial to reduce non-specific binding of antibodies.

Stripping:

If you are concerned about antibody cross-reactivity or need to reuse the same sample for multiple primary antibodies, you might consider a stripping step. This can be done using gentle methods such as glycine buffer or more aggressive methods like using a commercial antibody stripping buffer.

However, stripping can potentially remove or damage the mRNA signal, so it’s often better to plan your experiments to avoid the need for stripping.

Protocol Tips:

Sequential Detection: Start with HCR to detect mRNA, then proceed with IF for protein detection. This order is typically chosen because the HCR reaction conditions are less harsh and less likely to affect the subsequent IF steps.

Probe Design: Make sure your HCR probes are designed to minimize cross-reactivity with cellular proteins or other RNA species.

Antibody Validation: Use validated primary and secondary antibodies for IF. Ensure that the fluorophores used for HCR and IF are compatible and can be distinguished by your imaging system.

Controls: Include appropriate positive and negative controls for both HCR and IF to validate your assay.

Imaging: Use a confocal microscope or a microscope equipped with deconvolution software to minimize bleed-through between channels and to accurately localize signals.

References:

Choi, H., et al. (2018). “Simultaneous Detection of mRNA and Protein in Single Cells with Hybridization Chain Reaction and Immunofluorescence.” Cell Chemical Biology, 25(7), 950-958.

Stavis, C., et al. (2018). “Simultaneous Multiplexed Detection of RNA and Proteins in Single Cells.” Cell Reports, 25(1), 63-71.

Remember that protocols can vary greatly depending on the specific cell type, antibodies, and probes used, so you may need to adjust these recommendations to fit your experimental setup. Always start with positive controls to optimize each step before applying it to your samples.

Hey Buse, you could also use dPCR to detect CDs and other markers on the EV surface. I guess that is easier compared to staining, and you will not need too much antibody for it.

You can find the application note for it here: https://www.actome.de/downloads/Actome_EV_Quantification_App_Note.pdf

I found a way to achieve better staining. I used saponin 0.1% PBS instead of triton.

Adding 3% normal goat serum and 2% bovine serum albumin in 1xPBS 0.1% saponin also gives beautifully stained cells.

Glutaraldehyde is also horrendously fluorescent if you are using Immunofluorescence. Fixation with solvents will likely remove membrane proteins along with lipids. If the membrane proteins have an external epitope try staining live cells. Note that IF methods also typically include detergents to that may also remove membrane proteins.

Hi Mahdis Taghavibayat

Performing immunofluorescence (IF) on paraffin-embedded tissues does indeed increase the chances of encountering autofluorescence compared to frozen tissues. Here's why:

* Formalin fixation: Formalin, a common fixative used for paraffin embedding, can induce autofluorescence in tissues. It cross-links proteins, leading to the formation of fluorescent adducts that emit light when excited with the laser used in IF microscopy.

* Autofluorescent biomolecules: Paraffin itself can exhibit autofluorescence. Additionally, certain biomolecules naturally present in tissues, such as lipofuscin (wear-and-tear pigment), collagen, elastin, and red blood cells, can also increase autofluoresce.

The age of paraffin-embedded tissues can indeed affect the results of immunofluorescence (IF) and immunohistochemistry (IHC) in several ways.

* Antigen degradation: Over time, proteins and other target molecules (antigens) can degrade in paraffin-embedded tissues. This degradation reduces the number of available targets for antibodies to bind to, leading to weaker staining intensity and potentially reduced sensitivity of the technique.

* Reduced antigenicity: Formalin fixation, a key step in paraffin embedding, can mask some antigenic epitopes (binding sites) on proteins. Over time, these masked epitopes might become permanently unavailable for antibody binding, further compromising the staining results.

* Increased autofluorescence: Older tissues might exhibit higher levels of autofluorescence due to factors like further cross-linking of proteins by formalin and accumulation of autofluorescent biomolecules. This autofluorescence can mask the true signal from your target protein.

Please find the references attached.

* https://wellcomeopenresearch.org/articles/2-79

*

Thanks,

Personally I've not had to mount really thick sections, but I've seen other people make a shallow well on the slide by gluing coverslips to the slide. This might help keep the coverslip flat.

The appropriate stabilizing buffer for conjugating quantum dots (QDs) to antibodies is critical for ensuring the stability and functionality of both the QDs and the antibodies. The buffer utilized is determined by a number of factors, including the type of QDs, the conjugation chemistry, and the antibodies used. However, some general rules can help you choose an adequate buffer:

Example Buffer Composition

A commonly used stabilizing buffer for conjugation might look like this:

Conjugation Chemistry Considerations

Final Tips

Yes, confocal is preferred but I think you could use any type of microscope since they are slides and you can visualize slides both from the back and front, you should also try optimizing your staining protocol or maybe since we are talking about immunofluorescence, you could use another endothelial marker and see if you have a better signal. You can also try taking multiple photos at different focuses, however i find it extremely hard to do a 3D reconstruction of tubes, it also requires experience with image analysis and coding.

1) PFA is a very poor fixative if you want intact membranes. I'd try using just glutaraldehyde or glutaraldehyde and PFA mixed.

2) You must ensure that there are no large osmotic differences between the cells you will fix and the fixative you use (osmotic gradients).

Hi Ziqi Liu

If you are looking for a dye that stains lysosomes, lysotracker could be used along with required antibodies to stain the lysosomal markers along with lysotracker.

Here is a link to the dye website:

Hope it helps,

Thanks,

Malcolm Nobre, John Hardy Lockhart , Thank you so much

can you kindly show the graph? That will aid understanding of the question.

Hi Emily, You're correct that it's crucial to account for the area differentials between samples when quantifying signal from immunohistochemical (IHC) staining. As you've suggested, one approach to deal with this is to normalize the total signal to the total area of the sample, which essentially gives you an average signal intensity per unit area.

Here is a generalized step-by-step method using ImageJ (Fiji):

Repeat this process for each image, ensuring that you're using consistent settings throughout. By using this method, you're essentially determining the average intensity of the signal per unit area in each section. This can then be compared between sections, even if they are of different sizes. This is a simple and effective way to standardize your measurements across different sections.

Hi Carsten,

thank you for the suggestion!

I will discuss this with my supervisor.

Best regards,

Jan

Thank you so much. I will try this.

You could try blocking in donkey serum instead of BSA. Previously when I would do IHC or IF I used 5% goat serum in TBST as a blocking buffer when using secondary antibodies raised in goat. I have seen the percentages used vary between 3 and 10%. I used this buffer to dilute my primary and secondary antibodies as well.

There could be a number of reasons for weak or absent signals in immunohistochemistry, including:

Hi,

For the green, bright, fiber-like staining, it kinda looks like blood vessels...Red blood cells are highly autofluorescent and you might detect them if they were not completely washed during perfusion (the mice were perfused, right?). Did you check if you see this staining even before doing the immunofluoresence?

Also, which kind of primary/secondary antibodies are you using? Do you have a "secondary-only" control to check if they are causing such background?

Hi Ot Massafret,

Spontaneous differentiation is a rather chaotic process which can lead to several intermediate states. Is this occurring in vivo or in vitro? I’d suggest you let that differentiation run for longer and see if you still see those mixed/intermediate phenotypes.

Please check abcam company it might be useful for your experiment

Positive control

If you know your secondary works then it's looking like a problem with detecting your antigen. Your fixation is fairly standard but have you tried anything else like methanol or acetone? What retrieval steps are you using? If it's heat then maybe you need to try an enzyme instead such tryspin or pepsin (although these can damage antigens!). Have you got a control sample expressing your antigen such as a cell line etc that you can check your primary works on? Would give you confidence in the antibody and you could narrow it down to fixation/retrieval. Not sure if that helps!

generally, use a negative control with only secondary antibody in the whole staining process to ensure that this is actually a false positive stainings.

did you do this? furthermore, try to use different concentrations (dilutions of primary and secondary a.b.). that migght help too.

ensure that you use blocking solution and serum of respected host of primary a.b. to block.

lemme know!

silke

Staining methodology can influence uneven staining results. When applied directly to the material, first contact material is likely to take up the stain more intently. I do not know your staining protocols, but if a dilution of the sample is possible, such will encourage a more even staining result. A re-concentration of the material may be possible, post staining.

Narumi Uno Thank you for your suggestions.

Iskander Madhi I have not stained them yet, it is GFP after transfection. Thank you for your recommendations it would of help in the next step of my experiment.

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

Thanks for the advice. We make sure that the PFA is clear before filtration, and then we adjust the pH.

What we don't understand is that our PFA was working (using the same recipe as describe above) until end of 2021, and since then it doesn't fix anymore... We changed the PBS, the PFA, the pump, the flow rate, even the dH2O, and it is the same result.

Hi Dane,

Maybe you have already found the answer to your question, but what I read from your question can be answered if we take one aspect in consideration. Genomic Instability. Cancer cells are known to bear this as one of the causal features. With genomic instability, yH2AX occurance is normal. More the growth, more unstable the genome is. Before you come to a conclusion with this aspect, I would ask you to do a titration assay.

Grow the cells in 2.5% FBS, 5% FBS, 7.5% FBS and 10% FBS. Stain them. Lower FBS will lead to a slower growth. I expect you will see a gradient in the quantity of yHH2AX.

Somnath

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:

CD56 will stain NK cells, but other immune cells as well, so it's not a specific marker. NKp46 is a more specific marker for NK cells. Not sure you'll be able to find an IHC antibody for all MHC-I versions, and if you did it would probably stain all of the cells in the sample, so not sure what you are trying to do with that combo.

My ex-colleague used this trick to maximize binding of primary antibodies to a target, which presented in a very scarce amount - she just simply incubated slides at +2 - 8*C overnight. But I don't remember if it was FFPE samples or "wet" slices from cryotome.

Overall, PRNT has been reliable and a mainstay for virology and vaccine development. Here are the limitations I've found personally:

1) The throughput of the assay is somewhat limited due to the necessity of plaque counting as opposed to an automated read-out of reporter gene activity. I've really only done manual counts, and it looks to be somewhat difficult to automate the process. There are scanners with software for counting, but the visibility of the plaques and contrast with the uninfected monolayer can be limiting and the software may not give accurate counts. Related to this is the need to have the plaque size of the target virus being uniform so that counting is accurate.

One workaround is to examine the dilution series of each antibody and count only the two wells that bracket the endpoint, such as the 50% reduction in plaque counts. That can often be done in a hurry or to get preliminary data around the endpoint titer, but you lose a lot of information that way.

Another caveat for plaque counting either by human or machine is the necessity for cell monolayers to be uniform and complete enough in each well to get accurate plaque counts. If the cells do not form a 100% uniform monolayer by the end of the plaque growth against which the plaques can be counted, then it gets tricky, especially if the plaques/monolayers are stained with a dye rather than by immunochemical staining which may be better at showing the infected cells on a bad monolayer but is more time intensive.

2) The biosafety containment needs to be commensurate with the target virus biosafety. Often, researchers use the replicating virus of interest in the PRNT as opposed to a pseudovirus or chimeric virus which can be made to have lower biosafety requirements but will often have a reporter transgene if one is going to all the trouble of constructing these.

3) For some viruses that are more difficult to neutralize than others, it can be a little tricky to normalize the input virus between assays. This is more of a problem with the relationship between virus and neutralizing antibody concentrations, but practically speaking, inherent variability of the virus input can alter the results up or down depending on how much virus ended up in that assay. That can create more work in trying to put a large dataset together and have the assay data from each run match each other, say by the use of the results of a standard serum/antibody used in each assay.

4) One meta-problem that I've encountered is not the assay's fault, but rather the design of the assay with respect to the target cell. Viruses can enter the cells of various cell types in various tissues by different mechanisms, so the use of a non-physiological cell type for the neutralization assay can give nice, consistent results, but results that aren't useful going forward. For example, the human cytomegalovirus (HCMV) field for years employed primary human foreskin fibroblasts as the cells in the assay for which the neutralization of virus infection by antibodies was measured. This use is/was due to the very limited range of cell types that are available for permissivity of this virus for replication and plaque formation, so the cell type used is one of convenience (which itself is relative since these cells have limited passages). HCMV vaccine development experiments used these cells, and it wasn't for decades was it discovered that viral entry into endothelial cells is a more accurate assessment of neutralizing antibody responses compared to blocking entry/infection in a fibroblast. Thus, the fibroblast data wasn't particularly enlightening in the context of neutralization mediated protection. The cell type in the assay should be scrutinized when embarking on PRNT assays rather than treating them like another reagent in the assay.

I would recommend that you get the FIJI distribution of imageJ (https://imagej.net/software/fiji/) which includes a number of useful plugins for fluorescence microscopy analysis, including "Coloc2".

An overview of colocalization analays can be found here: https://imagej.net/imaging/colocalization-analysis#methods-of-colocalization-analysis

And directions for using the Coloc2 plugin can be found here: https://imagej.net/plugins/coloc-2#how-to-use-coloc-2

Hi Maynak Chakraborty ,

First remark: the dilution of your secondary antibody seems a bit high. It depends of course on the concentration of the stock solution, but I generally use 1:500 - 1:1000 dilution factor for most secondary antibodies.

Two questions now: 1) What is the species of your corneal endothelium tissue? 2) Do you use something as a blocking solution, like BSA or serum?

Hi Rafał Watrowski . Adity Pore was just asking why markers for the hypothetical EMT do not make sense in the 2 cell lines she is looking at.

I agree. The EMT is just a hand waving explanation of how epithelial cells, that are normally sedentary, suddenly become motile and invasive when they are cancerous.

Not at the relatively low concentration of these ions in this buffer. You wold need several M to possible have any adverse affect. Worth doing the incubation at room temperature or 37c to be sure. Reserve 4c for overnight incubation.

Vipul Batra

BSA might be the culprit provided that your secondary Ab is either anti-bovine IgG or anti-goat IgG or anti-sheep IgG. This is because BSA contain bovine IgG and above mentioned antibodies will react strongly with same (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3216515/).

Therefore, the use of BSA as a blocking agent may actually increase background staining. Some methods known to reduce non specific binding include using non-ionic detergents (such as 0.1% Tween 20) in PBS for washing steps.

Moreover, I have used up to 3% BSA in blocking buffer and it has worked in my case.

Fa Ro , that does make this quite a bit more complicated.

Fluorescently-conjugated small peptides (i.e. phalloidin) can be used to visualize some intracellular targets in live cells (actin filaments in the case of phalloidin), but this does not apply to most proteins. Unfortunately the size of normal antibodies prevents their entry into live cells, but a few different approaches for labeling intracellular targets in live cells (e.g. quantum dots and nanobodies) have been published. Unfortunately, none of these systems are in widespread use.

The most common approach for live cell visualization of a specific protein is constructing a fluorescent fusion protein. This can be done with transient transfection or through generation of stable cell lines. However, there is no guarantee that the fusion proteins will behave in the same manner as the wild-type due to steric hindrance.

One group has already reported a GFP-Eomes fusion protein in mice: . They used TALENs to insert the GFP into the first exon of Eomes, but you should be able to reconstruct this in a plasmid.

After separation your image into colours, you can enhance brightness & resolution with two important tools.

First : Image --> Adjust --> Brightness/ Contrast

Second : Image --> Adjust --> Window level

The final results have attached

With best Regards

You can permeabilize fixed cells with 10 min incubation with 0.1-0.5% Triton X then after washing and liquid removal you can put the semi-dry slides into -80 for indefinite storage. The permeabilization will allow the water to leave the cells without rupturing the membranes during freezing. I did staining on the cells fixed that way after 3 years of storage without any issues.

Overall, your procedure looks good. I have seen the drying procedure described before, but I have never personally done this. Leaving your slides out at RT doesn't seem to be a good idea. Some of the artifact you are seeing in your images (IMO) looks like artifact from the drying. Have you ever tried skipping the RT drying and moving your tissues directly to a -20C Freezer? Perhaps for an hour minimum? Drying tissue out is usually the enemy of all IHC. Also, you may want to consider .1% TritonX100 in your blocking and staining steps to increase antibody penetrance. Additionally, I usually use 3% BSA in all of my staining and blocking steps (1hr minimum at RT), including with neural tissue (seems to work more effectively than serum of the same species of secondaries). I wash with the .1% Triton (3 times for 5 min) and finish with a wash with PBS.

Another possibility is that the tissue is overfixed, which will also be detrimental to your staining. Be sure to wash the fixed tissue several times. Also, are you flash freezing in OCT?

I'm taking a guess, possibly if you have a positive control for membrane expression (transmembrane protein), say in Green color, and PI3Kbeta, Red color. Process growth factor (- neg cells, possibly serum starved cells) vs growth factor (+, pos cells). Fix with formaldehyde, permeabilize cells, stain and compare Red fluorescence/green fluorescence in - vs + cells. It should localize to the ruffles as you found. Excess ratio in +cells over -cells should give you the signal you're looking for. Image with confocal microscopy. Ruffles are a blessing in disguise.

Hi Cody,

There could be a few different things going on. Since the tissue is cutting well, you likely have a good fix and they have sat in sucrose for enough time.

From the looks of your picture it seems you should just add a little PBS to the slice so that it kind of floats in the liquid on top of the slide. That should give you enough room to use a brush and make sure the tissue is flat. Then, while gently holding the slice in place with the brush, angle the slide so that the liquid begins to run down the slide, causing the slice to 'sit' down on to the slide again. There should be enough residual liquid to allow you to make some minor adjustments to get your tissue perfectly flat on to the slide and position it where you want.

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

Hi Meara,

I think the easiest way is to use antibody to label endogenous reporter, like mCherry. I used GFP antibody to label endogenous reporter in paraffin section during immunofluorescence, and the staining is good and specific. Hope it will help you.

Best,

Chenglei

Hi Nam,

The short answer to your question is - no :(

To my best knowledge and based on my experience CD31can be detected on both live and dead cells, and even on isolated cell membranes. One may find some insignificant changes in CD31 levels in dead cells, but it is not a clear cut between the healthy and injured/dead cells.

Best

Gedas

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.

Dear Chaimaa Benlabsir

check the following link

it my be helpful

Dear Shahid,

Typically, polyclonal antibodies (pAb) contain a large number of different individual antibodies, which all differ somehow in affinity, selectivity and target structure. Compared to a monoclonal antibody (mAb), which may only show affinity to a certain target epitope of the antigen, polyclonal antibodies tend to target all available epitopes on the antigen. Therefore, depending on the used immunogen and the size of the antigen, the polyclonal antibodies are likely to bind all accessible epitopes.

As already pointed out by Wolfgang Schechinger, the mass is not affected by the binding site.

Regards Martin

hello, why don't you try to directly cytospin the cells?

We have done it before and we had very nice images.

After thawing of cells, we wash with PBS, then fix with methanol (ice chilled at -20) for 5minutes or PFA, then wash PBS, then cytospin the cells on slides. From there you can proceed with permeabilization and the rest steps of your IF protocol.

best of luck

Dear adity,

I think that you cannot decide to use only one instead of another. given that EMT is a pretty complex event you should decide according combinations of them.

for example: Twist-1 and Zeb-1 are two of the main four transcription factors therefore if you choose them. I would include also E-cadherin ( epithelial) and N-cadherin (mesenchymal) in order to verify the cadherins switch typical of this event.

moreover, assuming that tumors with increased activated pAKT and pERK together with decreased E-cadherin expression are destined to undergo EMT I would also include pAKT and pERK to have a more complete and wide view of the process.

last but not least: vimentin and cytokeratin in order to control if you'll have modifications of the intermediate filaments. please first verify which type of keratin your tumor specifically express in order to be more precise.

hope this can help

" Vimentin is a cytoskeleton intermediate filament protein present in cells of mesenchymal origin; this includes leukocytes, endothelial cells, and smooth muscle cells."

If I were to need the samples you require and were doing hypothalamic work, I would strive for separate samples for these purposes. However, if you are limited by finances, time, etc. and you are forced to take this approach, I would suggest alternating which side you keep for each purpose and using that as a factor in your analyses. For example, animal 1: left side IHC, right side western; animal 2: left side western, right side IHC, and so on. You can then determine if using left vs. right makes a difference in your particular assay/hypothesis using statistical analyses. It might make no difference at all but you don't want to inflate your chance of Type II error by just randomly selecting given what you know (or sometimes don't know) about lateralization of brain structures.

In addition to the above answer (by Suckert Theresa ), I have used Eriochrome black T, it works really well for skin and oral FFPE samples.

you may check this link for a related paper:

Please also have a look at the following RG links.

There are some good options out there.

Here's a good description of the concepts, and it lists a few plugins.

https://imagej.net/Colocalization_Analysis

I read this paper a while back but I haven't tried the plugin.

https://www.nature.com/articles/s41598-018-33592-8

You may need to do this with install: https://imagej.net/Following_an_update_site#Add_update_sites Giving the URL: http://sites.imagej.net/EzColocalization/

This ResearchGate thread is also pretty good: https://www.researchgate.net/post/How_to_quantify_colocalization_of_dual-color_immunofluorescence_image

There are also several videos on YouTube that take you step-by-step.

Have a very specific type of output you are looking for. For example, plugins like Coloc 2 don't give you information about how well your images/channels align with each other. Whereas, EZcolocalization may give you more of a per-cell-colocalization.

I hope that helps. Good luck!

- Melissa

Use OPAL kits from Akoya Biosciences and you can use any antibody (raised in any animal).https://www.akoyabio.com/phenopticstm/assay-kits-and-reagents

Kruskal-Wallis is fine IMO, I did that. I was interested in other options as well. Thanks for the input!

You might be able to couple the rhodamine carboxyl to an alginate hydroxyl with a water soluble diimide coupling reagent.

Or use a different dye....

hi paper,

you may coupling your macrophage staining with anti myosin heavy chain or perilipin immunostaining to mark fibers or adipocytes, respectively. Alternatively, you can perform a hematoxylin and eosin stain. In this case, adipocytes appear as white vacuolar unstained cells while fibers will be stained in red

Hi Chrisopher,

I had the same problem with a couple of probes that worked separately but not together. I solved the problem by increasing the dithiothreitol (DTT) concentration in the hybridization mix up to 1M. Maybe this can help...

The best program is Image-J software.

This paper is an excellent summary of DNA damage foci by Kai Rothkamm https://www.ncbi.nlm.nih.gov/pubmed/?term=foci%2C+meaning+and+significance%2C+rothkamm

Steve

To me this looks like blood cells. The reason why you are getting an overlap with DAPI could be just leaking of the green channel to the blue channel in the microscope (I think you are not using confocal, right?)

The sample preparation seems quite bad, there seems to be overlapped slices of tissue and uneven surface (the blurred areas). If the general quality is that bad, perhaps that's the reason for so many blood cells present. Can you get another sample?

Also, 1:400 could be a bit too concentrated... You know what? Try it without any antibodies, just DAPI. Do everything the same way you are doing until the antibody step, then skip both antibodies (no incubation at all) and go directly to mounting and imaging. If you will still get the signal, then it's blood cells for sure.

Hi Federico,

I don't know about HIF1a in mammalian cells, but in plants the HIF1a orthologue (RAP2.12), is sequestered at the plasma membrane during normoxic conditions to prevent N-degron mediated proteolysis: https://www.nature.com/articles/nature10536. Only when hypoxia occurs these orthologues migrate to the nucleus.

Again, I don't know if such a mechanism exists in mammals but if it does, this could explain why you see signal at the PM.

All the best,

Sjon

Hi there,

It depends on your cells, but we usually fixed with 4% PFA for 20-30 min.

Afterwards, you can remove the PFA-solution (just leave a few drops of PFA in the well-plates, it inhibits bacterial growth) and add PBS to the coverslips. If you close the well-plates with parafilm to avoid drying, you can store the cells for several weeks at 4°C and use them for staining at later time-points.

Best,

Sebastian

I do not think you need a special protocol. Use 10-20ul of antifade solution depending upon the slide size where cells are and cover with a cover slip. Be sure to avoid air bubbles. Keep slides in dark area on a a horizontal slide organizer. You can keep in refrigerator for several days or you can see immediately under microscope. You may or may not use nail polish around the rim of coverslip to avoid slippage of material outside. That is subjective. Keep antifade solution wrapped and at 4C

An antibody that does not work well in Western blots may be one that recognizes a structural epitope rather than a contiguous sequence in the protein. You still will have to prove that it recognizes the correct antigen with good affinity and specificity, but will need to use methods that leave the protein intact. Depending on the properties of your antigen, you could, for example, test whether your antibody is able to pull down a protein of the expected size from a cell extract, e.g. after gentle solubilization with detergent if you are looking for a membrane protein. If you have the purified antigen, you can also test your antibody in an ELISA setting.

Hello Rinki Di,

Thanks a lot for such a detailed explanation. I believe these tips will definitely help me in processing the brains in the near future.

And you are absolutely right about using the vibratome. I had very good results with vibratome in regard to sectioning in the past. Unfortunately, as for now I have already used cryostat to process the brains.

Thank you for your time and assistance.

Hi all,

First off, thank you all for your replies. Unfortunately, I do also see the high background when exciting with at 470nm and 590nm (I have included some 470nm images). Even with the reduced alexafluor 488 florescence in TDE, I was not expecting such high signal to noise ratio.

I was hoping to move up to 30-50um sections, which is why I have tried TDE as a mountant.

I have not tried the slowfade, but I am going to try prolong glass soon, and ill see how that goes.

great, thank you. Somewhere in the files we have data from a fluorescently tagged secondary AB, and I'll see if I can find the relative brightnesses of these QD's and that fluorophore.

I'll check out biosynth as well, thanks