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Immunofluorescence - Immunofluorescence - Science method

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Could anyone help me to figure out the cause of the stains in my immunofluorescence preparations?
I have been experiencing two different stains. First one is the bright striations that look like ir-fibres. The second one looks pretty much like a snowflake trapped under the tissue section. Aside from bad appearance it reflects the fluorescence light so that the surrounding cells can not be photographed properly.
We apply free-floating immunofluorescence protocol in frozen brain sections in our laboratory.
We applied the citrate protocol before the primary incubation in immunofluorescence staining then respectively rinsed the brain sections 3x5, 2x20 min with PBS. We incubated the brain sections in blocking buffer (%10 NHS+Immunobuffer) for 2h at 4C and thereafter incubated the sections in primary antibody at RT for overnight on a orbital shaker. We rinsed the section with PBS than performed the secondary incubation for 3h at room temperature. After all sections have dried we used fluoromount on the slide for mounting. Then we saw like ir-fibres and a snowflake like images.
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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?
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We are analysing the expression of Afp (endoderm marker), Sma (mesoderm) and Tuj1 (ectoderm) by mESCs that have been cultured in spontaneous differentiation conditions for 10 days with immunofluorescence staining, and we are detecting tha signal for more than one marker on several cells at the same time, i.e. we observe cells that are positive for Afp and for Tuj1. Is it possible that a single cell expresses more than one of these markers at once?
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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.
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Hi everyone,
I'm trying to do IHC staining with kidney tissue (Paraffin-Embedded tissue) with anti histone H3 (Abcam, ab503).
However, I could not find the positive control for the tissue.
In fact, the company only recommends PepArr, IP, ICC, WB; not IHC staining, that's why they only suggest positive control for WB is "Mouse and rat brain tissue lysates"
There are quite a lot manuscripts using this antibody for IHC staining, both Chromogenic and fluorescent IHC staining; but they don't mention about the tissue positive control.
I could not find any information why they suggest mouse and rat brain for WB control, I wonder if I could use that for tissue control of IHC staining.
If anyone has any experiences with this experiment, please share with me.
Thank you so much.
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Please check abcam company it might be useful for your experiment
Positive control
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We have been trying to visualize endothelial cells in heart sections, but it has failed every time. We have tried two different Vegf primaries and a Vegf R2 primary that a colleague has used successfully many times before. Our secondary antibodies (Alexa Fluor 594) work with other primaries (vimentin and Adamts2), but do not fluoresce with any of our EC primaries. We tried the Invitrogen Tyramide Boost Kits, both the streptavidin and HRO versions. For the EC slides the tyramide kits were entirely unspecific, but with our vimentin primary, they worked fine.
We have tested all of our secondary antibodies, the boost kits, and our protocol, and they all work on our cardiomyocyte and fibroblast markers, and specific gene primaries. It's only when we use our endothelial markers that they fail.
Any thoughts on what is going wrong or what we could change?
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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!
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I have a problem concerning my liver fluorescence staining: one primary antibody, GARP, in the attached picture in red, is causing these weird stainings on and around the nucleus. Does anybody happen to know how to improve my staining and get rid of these false positive stains?
Or general advice for liver staining, as the autofluorescence in liver tissue is always really high.
We are cooking our samples in citrate ph6 buffer, and using Avidin+Biotin as well as Tris,TWEEN20 + 5% BSA for blocking. In order to attach the fluorescence dye we are using Dylight 550.
I would very much appreciate any advice!
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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
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Hello everyone,
I am currently staining formalin-fixed paraffin-embedded skin samples with Opal IHC. All the antibodies for intracellular and cytoplasmic markers have stained well across all the samples except for DAPI. There is strong staining in a couple of samples but the rest of the samples have weak staining, where DAPI does not stain all the nuclei present. H&E staining of the same sections also show nuclei.
For these samples I have carried out deparaffinization with xylene and ethanol (100%, 90%, 70%, 50%), blocked with 3% H202 and performed antigen retrieval with citrate buffer (pH6) four times. All the samples were mounted with ProLong Gold and all the washing was performed with TBST. I have tried NucBlue Fixed Cell ReadyProbes DAPI using 2 drops in 1mL TBST and both a 1:500 and 1:1000 dilution of DAPI dissolved from lyophilized powder with similar results. For both DAPI used, I have left DAPI on for 30 minutes.
Does anyone why DAPI isn't staining equally across all my skin samples and if this is issue with DAPI or the actual samples?
Thank you.
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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.
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Hi dear friends,
I am working on Parkinson model. I transfected SH-SY5Y cells with topo isomerase IIB by lipofectamine 3000 reagent. I took some photos today but their backgrounds were too noisy. Notably, I took them by different channels, but I have got the same result. Worth to mention that we are sing ZEISS.
My question is what can I do to solve this problem?
I have added photos below.
Thanks in advance.
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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.
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Hello everybody,
I am doing immunocytochemistry staining for splenic B cells from WT and Knockout mice for one protein. By FACS analysis, it's clear that the signal for the knockout protein is nearly absent in Knockout cells, but when I do confocal microscopy experiment for the same cells, the signal seems to be the same in knockout as in WT. Does anyone have an explanation or can recommend what to do?
P.S: I use the same primary antibody for IF and ICC and No background for the secondary antibody in the control.
Thank you!
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Thank you Alibek, I will do it and I hope it will work :)
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I prepare the PFA 4% following the recipe of 40g in 1L of PBS 0.1M. The pH is neutral (between 7.2 and 7.4), the solution doesn't boil (I keep it under 60°C).
However, when we perfuse our rats, the PFA doesn't fix them at all, they remain very soft.
Does someone has any clue on why the PFA is not fixing ? We tried to change the provider of the PFA powder, the PBS, nothing worked...
Thanks in advance for your help.
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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.
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DNA Damage Question
I am performing Phospho-yH2AX immunofluorescent staining of cells treated with either 0.1% DMSO or a drug inhibitor in order to measure DNA damage. Unexpectedly, I am observing high levels of signal in the DMSO treated cells. The cells are grown on Chambered Cell Culture Slides from CellTreat. It does not appear to be cell type specific given I have observed this with 5 breast cancer cell lines and 2 prostate cancer lines. I am using the CST Phospho-Histone H2A.X (Ser139) (20E3) Rabbit mAb #9718 antibody in Normal Donkey Serum at the recommend 1:400 dilution, so I don't believe its non-specific signal either. Has anyone encountered this issue before or know of any alternative explanations.
For reference I have attached an image below where H2AX signal is shown in pink these are completely untreated suggesting it is not an effect of DMSO
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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
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Hey!
I have a FRET system between the inner and outer nuclear membrane, and I was wondering if I stained with DAPI or any other fluorescent antibody if that would mess up my FRET readout? As of now I am using acceptor photo bleach and I am working with neuroblastoma cells (Be2-c). I can do either fixed or live cell imaging, depending on which would work best if this even possible.
Thank you!
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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:
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I am working with patient-derived tumor samples and I would like to detect CD56/NCAM1+ cells as a marker for NK cells and MHC-I molecules (HLA-ABC) in the same slide.
Could you recommend chromogenic vs fluorescent detection and, if possible, specific antibodies to build my panel?
Many thanks
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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.
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Hi, I am Joe and currently, I am working in a neuroscience lab and focusing on the adrenergic beta and alpha receptors. And I have some technical problems with that, which in my experiment there is always low efficiency to stain the alpha 1and beta 2 receptor in mice brain. I have already tried several antibodies including the Almo, Bio, thermo antibodies
Therefore I would like to ask if there are any tips to increase the staining efficiency and do you guys have any recommendations of the antibodies for the Beta 2 and alpha1 receptor for mice? Thank you so much for your help.
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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.
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Hello there, i'm working on a differente approach for Plaque Reduction Assays, and i would like to ask you, professional working with this type of technique: which are the major problems found by you when working with PRNT?
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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.
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I have limited amount of ethanol and ample amount of methanol, I was wondering if it can be used to make gradients of alcohol for Histology processing(staining). I know methanol is not preferred because it is toxic in nature. but is the toxicity going to effect the formalin fixed tissue ? Is it going to effect antibody binding if i perform immunohistochemistry/IF on the sections. ?
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Interesting question. I think, you mention the main concerns why methanol is often not preferred over ethanol. I found this on one website: "Methanol This reagent is a good ethanol substitute but is rarely used because it is volatile, flammable and costly. Methanol tends to harden tissues more than ethanol and is a poor lipid solvent." https://tissuesampling.weebly.com/processing.html.
[see also "Methanol is very rarely used in routine processing due to its flammability, volatility, and cost.[3]" from https://jprsolutions.info/files/final-file-5da2c9a0c9f2a9.26858550.pdf
I am not sure methanol is really more expensive than histology grade ethanol, or denatured alcohol for histology ("Anhydrous reagent grade alcohol is a denatured form of ethyl alcohol, approved for sale under U.S. regulations (90 parts ethyl alcohol, 5 parts methyl alcohol, and 5 parts isopropyl alcohol). It is a preferred mixture for dehydrationg tissue samples prior to embedding in histology.") contains methano). I guess addition of methanol is mainly to make the alcohol useless for human consumption.
Based on all this, it seems feasible to switch to methanol for histology purposes. I am sure it will change the hardness of the tissue compared to ethanol but I have not tried, and I do not know whether it makes the tissue harder and more brittle or the opposite.
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Hi all,
I need to learn how to use ImageJ to do colocalization and quantification of my IHC mice brain slides.
I have mice brain sections that I stained with the microglial specific marker iba1, and costained it with M1 and M2 markers iNOS and Arginase-1, with DAPI as a counterstain.
So my slides are as follows:
DAPI
iNOS --> Alexa Flour 488
Iba1 --> Alexa Flour 594
And
DAPI
Arginase 1 --> AF488
Iba1 --> AF 594
I couldn't find a resource on how to quantify this, any help would be appreciated.
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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
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I am doing Immunofluorescence assay to detect the target protein expression level in corneal endothelium tissue. But, in the confocal microscope, I have found background signal in negative control ( no primary antibody, only secondary antibody) image. The pattern of the signal is also very different from the positive control. I have used Goat anti-Mouse Alexa 488 as a secondary antibody (1:250). If anybody has any suggestion on how to get rid of this problem, it would be appreciated. The data is attached below.
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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?
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Hi everyone, I'm running an in vitro experiment with rat astrocytes and meningeal fibroblasts and I'm looking for antibodies to stain both cell types and differentiate between the two. I use 4% PFA to fix my samples, but other methods are also welcome. Which antibody would you recommend for the fibroblasts? For the astrocytes I use GFAP.
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only a subset of fibroblasts express astrocyte markers, such as ALDH1L1, most of the fibroblasts express markers of oligodendrocyte precusor cells, such as NG2, Olig1, Olig2, PDGFRa. Even for OPC, you can also dissect them by new subtype markers, this is still ongoing for investigation.
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I tried staining MCF7 and MB231 for Vimentin and Cytokeratin antibodies, and surprisingly, I saw both cell lines showed both markers. According to the literature, MB231 is a mesenchymal type cell and MCF7 is luminal, so shouldn't it be that MB231 shows more cells positive for Vimentin as compared to MCF7?
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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.
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Hello,
I am trying to stain glioblastoma cells with an anti-CD56 antibody in PE. For adherence of the cells, I am using PBS with magnesium and calcium.
Almost every cell seems to express CD56, but I only obtain very weak signals in the PE channel. Even when I am adjusting the exposure time.
Could this be because of the PBS that I am using? I've read somewhere that ions might hinder antigen bindung of the antibody...
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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.
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I have raised antibodies against 3 Buffalo beta-defensin proteins in Rabbits. However, while doing ICC or WB, a signal is always there. I have diluted enough (1:2000 for ICC and 1:80000 for WB), blocked enough (5% BSA in WB and 1% BSA in ICC) but couldn't get rid of it. The buffalo defensins' sequences are very different from Rabbits' i.e. very low homology. What could be the cause of this positive signal? The bands in western and binding pattern in ICC are similar for pre-immune sera and immunized sera. I have tried sera as well as purified antibodies from those sera. P.S. Could BSA be the culprit? Dont tell me that please
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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.
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I want to visualize EOMES, a transcription factor, in CD8 T cells. I am planning to perform an immunofluorescence test and I was wondering what kind of stain or chemicals I can use to enable the visualization of internal markers of a cell.
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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.
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Hello,
I am using RFP-GFP-LC3B for autophagy flux detection. After merge RGP-LC3 and GFP-LC3 result to a merged pic with yellow and red dots of LC3. I wonder how to seperately count red and yellow dots on the merged pic as the ref detached below?
many many thanks!
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In the end
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
Notice: You have to create a duplicate from the blue channel (Channel 3) before editing.
The final results have attached
With best Regards
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I fix blood cells on slides using 4% PFA for 20 mins at RT. I always keep an extra slide in storage, in case if I need to re-stain the slides. So far, I have been storing my slides at 4C in a petri dish with parafilm on top. The surface of the slide where the sample is, is covered with PBS (Attached is a picture). I have seen drying happening in 2 months if I don’t replenish the PBS on my slides. Is there a better way to store these slides for longer periods without drying or having to replenish? If I have to store these slides at -80 C, will I need to add some cryopreservatives like DMSO or these slides can be directly stored in -80 C? Please advise.
Thank you.
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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.
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I am using rat optic nerve tissue that was perfuse fixed, transfered to an azide solution after 24 hrs. frozen and cut at 14 microns on a cryostat.
General protocol
Dry at rt for 1hour
Tris Edta pH9 AR (because we have little to no staining if we dont)*
Pap pen, 3x PBS wash
20mins background sniper blocking (we have extensively tried various blocking and no blocking)
3x PBS wash
Primary antibody overnight at 4 degrees
3x wash
Secondary antibody at RT for 2 hours
The issue is that everything will be working and look good, then for absolutely no reason I can find the tissue seems to just absorb everything. I have optimised these antibodies individually and in combination before but sometimes Ill just get images like this and I cant work out why
The images are Cyclin D1 in 405
ASPA in 488
MyRF in 555
8OHDG in 647
It looks to me like the 8OHDG is the only antibody that appears to have worked, I have no idea why. Can anyone advise?
*We have tried with other AR solutions and without any AR
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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?
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I am currently trying to stain for PI3KBeta in MDA-MB-231 cells. My current issue is that 1) beta stains all over the cell and 2) there is an increase in signal at the edge of the cell and this increase in signal co-localizes with cellular ruffles (imaged in phase). My goal is to look at PI3Kbeta's recruitment to the plasma membrane in response to growth factor stimulation; however, an increase in staining at the edge of the cell that may be due to ruffling makes it very difficult to study this. Does anyone have any tips/experience/references that can help me analyze protein recruitment to the edge of the cell and reduce the effects of signal increase due to ruffling. Are there any techniques to keep the cell flat on the coverslip? thank you so much for your help!
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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.
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I am cutting rat brain sections at 20um at -20C. The tissue is cutting nicely out of the O.C.T solution, but I can not get a decent section to adhere to the slide.
I have included an image.
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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.
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Hi,
I'm having problems with the Recombinant Anti-MAP2 antibody from Abcam (Recombinant Anti-MAP2 antibody [EPR19691] (ab183830)) antibody on my frozen, 4%PFA perfused, mice brain tissue.
I was wondering if someone could elaborate better the note from the application part of the data sheet; Antigen retrieval: Heated citrate solution (10mM citrate PH 6.0 + 0.05% Tween-20).
To my knowledge AR is not used with frozen tissue, as it is to rough, and can damage the tissue, while also causing tissue detachment from the glass!?
Does someone have any experience with AR and frozen tissue?
I did try using 1/100 dilution, 0,2% triton X-100 and 4%PFA fixation as described in the images part of the datasheet: Immunohistochemical analysis of 4% paraformaldehyde-fixed, 0.2% Triton X-100 permeabilized frozen Mouse Cortex tissue labeling MAP2 with ab183830 at 1/100 dilution.
However I did not observed cytoplasmic staining of my neurons (image as uploaded for reference)!!
My protocol was the following:
4%PFA fixation, 15 min RT
3xPBS, 5min
10 min PBS+0.2% Triton x-100
Blocking solution (3%BSA, Without Triton x-100, 1h)
Anti-MAP2 1:100 dilution in 1%BSA (try both overnight and 2h RT)
3xPBS, 5min
AF-647 1:500, 30-45 min RT
3xPBS, 5 min
DAPI, 5 min
3x PBS, 5 min
dH2O, 5 min
Antifade and coverslip.
The antibody works excellent in IHC-P.
Thanks for the help,
Fran
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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
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I am trying to do antigen retrieval on mouse brain sections to stain for proliferative markers like MCM2, Ki67, PCNA etc on tissue that has mCherry conjugated to a transcription factor of interest to out lab. However, most AR techniques seem to kill the mCherry signal after staining. I have use many different heat induced AR methods with a variety of citrate buffers but any antigen retrieval that brings the tissue above boiling seems to kill the signal. What are some other options I may want to pursue?
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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
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Hi,
I am wondering if the immunofluorescent staining of CD31 antibodies can both mark endothelial cells and indicating their viability. In other words, if I perform immunofluorescent staining for CD31 on a sample of HUVEC, will the signal correlate with the cells' viability?
Your help and relevant references are highly appreciated.
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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
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I did a Western Blot for a protein that should only be present in hematopoetic cells. The WB shows a very clear picture, present in k562(hematopoetic) and not present in HeLa and 293T. However when using the exact same antibody for IF on HeLa cells we do see clear signal from our protein of interest (see picture). Can anyone provide insights to how this is happening? in controls without primary/secondary antibody we see no signal.
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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.
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My lab have some skin biopsies that were previously conserved in paraffine , and i wanted to know if it is possible to extract DNA or RNA from them.
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check the following link
it my be helpful
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Hi guys,
I m wondering to know my pAb binding sites to the antigen. I am not sure about my pAb's binding towards N terminal or C terminals of antigen. How I can check this? Also, does molecular weight change based on binding sites (N or C) of Abs?
Can anyone help me?
Thank you in advance!
Shahid
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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
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I am a physicist, so I know the techn not the biological applications. The question is straightforward. I am looking for a diagnosis that rely on fluorescent based cell imaging where the intensity of the expressed marker is the key part of the diagnosis. Thank you !
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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.
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Dear,
We have collected PBMC and frozen in nitrogen and store them at -80oC. I would like to ask whether I could do IF with these cells by attaching them to the coverslip. I found the protocol from this group:
and want to follow their method. However, I am not sure whether it works with frozen PBMC.
Thank you so much.
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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
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There are several EMT markers in cancer, such as vimentin, n-cadherin, SNAIL, TWIST, ZEB etc. How do you decide which one to use?
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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
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I am staining patient blood sample slides for identifying CTCs and I use the epithelial marker pan cytokeratin and mesenchymal marker Vimentin. I stain WBCs for CD45 marker. And I have seen that almost a big chunk (~60%) of my WBCs stain positive for vimentin . I am using the technique of immunofluorescence. Can someone please tell me the reason for this?
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" Vimentin is a cytoskeleton intermediate filament protein present in cells of mesenchymal origin; this includes leukocytes, endothelial cells, and smooth muscle cells."
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I am looking to complete western blot experiments and IHC experiments on the same cohort of mice.
Is it possible to Harvest the mouse and use the right hemisphere for IHC and the left for western blot?
I typically run simple tissue perfusion right after harvesting which runs perfusion on the entire tissue. Could I do this and still half the brain using each hemisphere for a different experiment.
We have done this previously only extracting the cerebellum but was much less invasive.
Thanks for any advice!
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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.
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I have been using conjugated antibodies for staining cancer cells. And I try preserving the fluorescence signals using Prolong antifade with Dapi. However, after 24-48 hours, the only signal I see is DAPI. All other signals such as FITC (cytokeratin) and TRITC (CD45) are lost/ barely visible. Is this a common problem with conjugated antibodies?
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In indirect immunofluorescence technique, multiple secondary antibodies will bind to the primary antibody used to bind the protein of interest. This allows us to get a better signal than just using a conjugated primary antibody. In your case, looks like the problem comes from photobleaching of FITC and TRITC ( I am assuming you were getting good fluorescence initially but lost after preservation). I think you can try alexa conjugated antibodies instead of regular FITC or TRITC. I have used alexa 488, alexa 647 and alexa 568 conjugated antibodies. I found them preserving fluorescence signal quite well.
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I used Fluorophore Conjugated primary antibody (Alexa-488, 555 and 647). Tissue FFPE samples were imaged before I stainded antibody and I found background fluorescence.
So, my question is "what is the best of reduce autofluorescence in FFPE tissue using between sudan black b , TrueVIEW® and TrueBlack® ?"
And If you any methods for reduce autofluorescence please tell me.
Thank you
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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:
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The situation is going worse and the health system in IRAQ is so poor to provide the simplest treatment needs of patients. What should we do in such a horrible situation?
Is there advice that should we follow to stay safe and healthy until they come up with the vaccine of COVID-19.
How to make our immunity stronger in this situation, what should we eat? what should we drink? what kind of medicine should we have to have at home?
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Hello,
Does anyone know of methods (preferably using Fiji) to assess quantitative data to analyze double positive cells. So I have figures from FITC/TRITC/DAPI channels and not all of my signals overlap and I would like to have a method to show that certain percentage of cells are overlapping.
Are there any plugins or other tools besides FIJI to assess that?
Any help would be greatly appreciated.
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There are some good options out there.
Here's a good description of the concepts, and it lists a few plugins.
I read this paper a while back but I haven't tried the plugin.
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
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Hi every body,
If the presence of a specific protein is not approved by doing IHC, can it be enough to conclud that this protein did not expressed in this sample ? Is it needed to do othere experiments such as western blotting?
If a protein differentiated or transdiffereencited to othere , how will change result of IHC?
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Hello, my lab is intending to try a triple immunoflourescence (IF) stain (anti-GR, anti-MR, and anti-cFos). We have been instructed by my lab tech to look for primary antibodies that are from different host species, otherwise there will be cross-binding. However, all the companies I have looked at only sell antibodies with host species rabbit or mouse. There is the odd goat/donkey/bovine host, but these are quite rare, and are not available for the specific antibody I am looking for.
Is it possible to do IF without having three distinct animal species hosts (i.e., all rabbit).
Thank you!
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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
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If we have multiple cell-types and multiple marker expressions labeled with a standardised immunhistochemistry (IHC) protocol which is the best correlation method to show a link between protein expressions in different cell types? If you apply a treatment can you use or is it better to use a different one?
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I am glad I could help you Sir! Kruskal-Wallis is a nice option.
Also I suggest to check your variables (if you measured multiple variables) with a PCA. It helps you to see the similarity of the tested groups (cell types in your case) and even to see how your measured variables correlate with each other.
For more info on PCA:
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Alginate is an anionic polymer and on the other hand Rhodamine B is a cationic dye. Theoretically these two should be boned by ionic bond. Does it happen? More suggestions and experience are greatly appreciated. Advanced thanks!!
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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....
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I am performing macrophage staining in the oral mucosa using an IFA protocol. There are oval-shaped structures on my scans that have background stain/autoflorescence. After doing some research, I found they are either cross-sections of skeletal muscle or adipose tissue. They sometimes appear ordered and clumped like they are skeletal muscle cross-sections, while other times they are more sporadic as if they would be adipose tissue. I am having difficulty identifying which is which. Do both tissue types usually autofloresce? If so, how can I differentiate between them while looking at my scans? Thank you!
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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
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Has anyone done FISH on cre-recombinased mice? I want to know if anyone has successfully been able to quench the autofluorescence from the those mice to successfully visualize the targeted genes? I have cre-mice crossed with ROSA26 tomato mice that I would like to do FISH on.
Any help or guidance would be greatly appreciated. Thank you!!!!
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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...
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Hi!
In our project we need to analyze a certain protein secretion by cell. We did a RT-qPCR but the time left is not enough for western blot. We had the immuno-stained fluorescent images of the samples and gonna use the images as a quantification of the secreted protein using ImageJ. I checked several algorithm online but was still not sure how to canonically present the quantification as a measurement of the target protein. Could somebody offer more information and give some papers that use quantification of fluorescence in image as measurement of practicle amount?
Thanks!
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Hi!
In my personal experience, I used the program Fiji to quantify the immunostainig of a particular protein. This program enables you to calculate the tissue area, the positive area and so on, thus permitting you to normalize the positive area on tissue area (if you have tissue sections like heart setions, you might subtract the not-tissue area from the total area of your picture, thus obtaining the real tissue area). This method, however, is very long, but you can use it especially if the number of the pictures that you want no analyze is not too high. I learnt this method at Professor Sironi's laboratory (University of Milan), so maybe you can find more information searching online Laura Castiglioni/Luigi Sironi papers.
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If I have more number of foci, does it represent nuclear damage or nuclear repair? Some cells show pan nuclear staining, does it mean the cell underwent apoptosis?
I am new to this area. Any help would be appreciated.
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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
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Hi all,
I am currently running trials of IHC using fluorescent labeled secondary antibodies to detect pyroptosis markers on rabbit tissue. I am seeing fluorescence in tissue treated without the primary antibody, where there should be none. This fluorescence is also exactly overlapping the DAPI stain. I am using paraffin embedded tissue, alexafluor488 secondary antibody, and my antibody species are all correct in terms of my host species. I have attached a picture of the type of stains I am getting. My secondary antibody is at a 1:400 dilution.
Any help would be much appreciated!
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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.
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Hello every body,
I've done Immunofluorescence on a canine histiocytic sarcoma cell line (DH82) and on the same line persistently infected with a virus for an oncolytic related project.
I've performed it both on FFPE cell pellet and on cell fixed on 96/well plate.
I've tried several markers for the project with same protocol and everything is good, the problem is with HIF-1ALPHA.
I've for sure cytoplasmic and/or nuclear coarsed area of positivity for HIF-ALPHA but both the cell type express frequently a strong MEMBRANEOUS pattern of staining, is it true or just aspecific?? (FFPE-red) (Well plate-green)
I've red that might be real, but anyway, why ?? to be honest I can not justify this localisation for a transcription factor
Would you suggest just to count only the nuclear and cytoplasmic positive cells as positive and to exclude the membraneous from the statistics?
Thank you very much for your precious help
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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
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I use 4 % PFA to fix my cells on a coverslip. How long should the cells be treated with PFA? Also, how should I maintain these fixed slides for longer periods? Are these slides stored with some liquid to avoid drying?
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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
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My cells are fixed on a coverslip and are stained with DAPI, Cytokeratin and CD45. What steps should I take after staining the cells?
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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
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I have a home-made affinity-purified antiboy. It works badly in WB but turn out to be great in IF. But we should test the specificity of the staining. However we don't find any mutant of this gene available. So my PI proposed that:"you can pre-incubate the antibody with the antigen, then use this to perform immuno-staining. If the observed staining pattern is gone, it would suggest that the antibody is specific.“ So how much antigen should I mix with the antibody? Does anyone has a proper protocol? We do the staining in how mount zebrfish brain issue(laval brain).
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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.
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Hello experienced people in IHC/IF,
I am doing an IF experiment to stain free-floating, 30 μm, mice hippocampal brain slices for a membranal protein. However, when I made sections using a cryostat, I found that my slices rolled up upon transfer to PBS. I used 1% PFA+1% sucroce in 0.1M PB for light perfusion of the brain (as per the protocol for my protein of interest). I embedded the brain in OCT before cutting and used the anti-roll glass while sectioning. The slices looked fine until I put them in PBS. I was wondering if anyone knows how to unroll them using any known alternative methods.
Thanks for your help/suggestions in advance.
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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.
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Hi All,
I have recently tried using TDE as an alternative mountant to mowiol to decrease z-axis aberrations for thicker sections. However, i find that the TDE creates really high levels of background florescence in my tissue sections.
I have included images of +/- DAPI stained TDE mounted sections, and a DAPI stained mowiol section as examples.
I was wondering if anyone may have had a similar experience?
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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.
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Using a fluorophore rather than a chromophore for ELISA detection seems like an obvious solution to increasing ELISA sensitivity, but I'm struggling to find work done in this area. Can anyone recommend a molecule sensitive to HRP/H2O2 that can give a bright fluorescence endpoint? We've tried fluorophore-labeled antibodies and saw no increase in sensitivity.
I have a flash-lamp sourced, monochromator-based fluorescence plate reader, not a laser-sourced one, so AlphaLISA is not a viable option.
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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
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I routinely functionally assess the iPSC derived neurons using calcium imaging with Fluo-4/AM (490nm/510nm). However, have not been successful at using those same neuronal dishes for carrying out ICC. The reason that I think could be are as follows:
  1. The calcium-sensitive dye fluo-4/AM has an emission at 510nm. As a reason why, there is a spectral overlap with my anti-bodies of interest.
  2. As I try and wash out the dye from the dishes, the neurons detach as a matter of fact. These neurons have been subjected to constant washing before and after the calcium dye-loading process. Moreover, these neurons are first stimulated with TTX following ionomycin and EGTA+TX are added to the dishes as internal controls.
Any suggestions or references or direction to a protocol would be of a great help!
Thanks!
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Hi all.
I have an application which requires direct staining whole blood sample with CD markers. Currently I'm having problems because the fluorescence signal from the stained samples is weak and sometimes I observe clumping of cells.
My protocol is
1. Aliquot 50ul freshly drawn whole blood in EDTA
2. Add Fluorescent conjugated CD45 antibody according to manufacturer's recommendation
3. incubate in dark for 20min
What I can do
1. add excessive CD45 antibodies
2. instead of adding antibody to whole blood, add antibodies into test tubes first then drop whole blood in
3. look for a very bright fluorophore as conjugates (any suggestions?)
Unfortunately I cannot fix the blood sample for this research.
Ultimately, I am looking for a way to make the stained cells as bright as possible, any suggestions on what I could do?
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Kevins answer above is the best way I think - and usually CD45 should not require any bright fluorochrome - it is the most expressed cell surface antigen on leukocytes - about 10% of all proteins if I remember it right. Your main problem is that the number of erythrocytes is so much higher than the leukocytes, so even if you collect more than 1 x 10^6 events it will be hard to spot the leukocytes - almost everything will be erythrocytes. Moreover, since you like to keep the cells unfixed you need to make sure that the you use a non-fixing lysis buffer - for example BD PharmLyse will not fix the cells, while BD FACS Lysing solution will fix them - you need to read the info carefully before deciding which to use. Essentially any solution based on Ammonium lysis works (can also be made yourself if you search the net). You can either wash the cells or analyse them directly with the lysis solution present (they will be rather diluted but it is OK if you have the time and not to much tubes). For data analysis it may be better to gate your cells on SSC/CD45 before SSC/FSC if you have a lot of debris (and use a CD45 threshold while collecting so that you do not even see the debris). You will then easily see your major populations (lymphocytes, monocytes and neutrofiles) - your gates can then be confirmed on SSC/FSC where the populations should also differ extensively. (Lymphocytes - low FSC, low SSC, high CD45; Monocytes - medium SSC, high FSC, high CD45; Neutrofiles - medium FSC, high SSC, low CD45).
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I am trying to see effect of some chemicals on brain receptors. I can only access small area even with 10X. Are there ways to access whole brain and see overall effect on all the areas.??
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Hi Rupak,
If you just want to see an overall effect on a specific staining, maybe confocal microscopy is not necessary. If you have access to an imaging facility, the may be equipped with a slide scanner such as Nano Zoomer.
This machine allows you to scan whole sections (such as rodent brains) with a good resolution, in brightfield or fluorescence. Of course, you won't have any resolution in z, but in your case, it's probably not that important.
Here is the website where you have more information about NanoZoomers : https://nanozoomer.hamamatsu.com/jp/en/product/search.html
Good luck,
Vincent
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- When I performed ICC work for treatment cells in translocation stress experiment in IBIDI 12 well removable chamber I got variant signal intensities ...
What is the reason behind cells stress without any kind of treatment!
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Thanks Udesh for participation
Actually Im using
- a removable 12 well silicone chamber for adherent cell lines and immunofluorescence staining...
  1. - Dimensions of wells (w x l x h) in mm 7.5 x 7.5 x 8
  2. - Volume per well 250 µl
  3. - Growth area per well 0.56 cm²
  4. Primary and secondary antibody diluted with blocking buffer (BSA;TRITON 100X;PBS ) to a final volume of 1.250 L
  5. I used to add 60 ul per well nut now I used 100 ul but with the same results even with high low confluent cultures...
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Hello! I am planning to do IF on adipocytes in cell culture but I have some questions. I will use 2 conjugated antibodies from different hosts and I can not be sure that there is any way to do double staining IF with conjugated primary from different hosts (I will not use seconder antibody). I found some informations, they had prepared a cocktail from antibodies but their antibodies are from same host. Can I use same way for antibodies belong to different origins? And my other question is one of my antibody will stain on a organelle (mitochondria), my other antibody will stain cell membrane. Should I use different protocols? Could you inform me about these questions?
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Shen-An Hwang Thank you so much for these valuable informations. I will pay attention your all knowledges. Have a nice day.
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I am trying out immunofluorescence on rabbit spleen but the results are not fruitful due to auto-fluorescence. I tried blocking with 5%BSA for an hour still persists. I also tried sudan black but could not see any signal, even in the positive control. Maybe i am doing it wrong.
Help from the experts in the field is much appreciated.
Thanks in advance.
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Dear Narendran,
Indeed autofluorescence can be a drag and some tissues are worse than other. I guess lots of autofluorescence in spleen comes from haemoglobin. If you have the chance to use confocal laser scanning microscope than the lambda scan over the whole wavelength range can help you determine which wavelengths emit the least autofluorescence and than just hope that your specific signal would be higher. This way you can define the signal treshold on the negative sample and without changing any acquisition settings collect the positive signal above the treshold from your samples.
Hope this helps. Good luck.
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In our lab, we freeze tissue samples from mice (embedded in OCT); we used to have a few problems with this method, using the standard plastic 15 mm cryomolds (frozen with dry ice & alocohol; or sometimes liquid nitrogen instead) -
1) It was very time-consuming. We would hold each individual cryomold over a beaker for several minutes - not only does this require a steady hand, it's slow and doesn't allow for multi-tasking. We freeze several samples a day, so it was taking up a lot of our time.
2) The frozen blocks can be difficult to extract from the mold.
3) Not enough space for labeling samples on the standard plastic cryomold.
4) Difficult to store samples in the mold; we would need to cover each with tinfoil before storing in the freezer (not optimal for protection of the frozen samples).
5) Samples would dry out in the freezer; we would need to wrap them with saran wrap to better preserve the mold prior to cutting.
I'm curious to see if anyone is having the same problems, and if there are any suggestions to improve the process?
For our own solution, we created a four-mold cryotray + box with platform - it lets us freeze four samples at the same time, hands-free. Each mold also has a lid for better protection/storage/hydration retention; and more space for labeling than a typical mold (see: www.sealnfreeze.com).
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Hi Dmitry:
I am not very sure about the 'dry out' problem in your experiment. But if it means many 'Swiss-cheese holes' inside your stained tissue slices, they were not caused by tissue drying in the fridge, but caused by ice crystals formed during your freezing process. To prevent this problem, tissues must be frozen as rapidly as possible. The EtOH-Dry ice bath did not provide a snap frozen environment (it's only ~-80 degree), and embedding tissues in big OCT chunk further slowed down your freezing speed (OCT is highly resistant to temp changes in surrounding environment), that's why you felt your freezing process was very slow.
My suggestions: 1. Use liquid nitrogen cooled isopentane to freeze tissues instead of EtOH-Dry ice bath. It would make your tissue frozen very rapidly and prevent forming big ice chunks inside tissue. 2. Do not embed tissues in OCT when you freeze tissue, but put tissues into liquid nitrogen pre-cooled isopentane directly. You need to determine the freezing time empirically: large tissues require more time to freeze but too much time in isopentane would cause tissue crack into pieces. You can store tissues in -80 fridge for short time period and embed tissues in OCT later. 3. When you embed tissues in OCT, set the temperature of
freezing microtome chamber to -6 to -8 degree (the freezing point of OCT in -10 degree) and use this temperature to pre-cool the OCT (Otherwise the room temperature OCT would thaw tissues and the ice crystals would be formed again by slow re-frezzing process). You also need to embed tissues inside the cold freezing microtome chamber. After embedding your tissues, transfer OCT chunks into a foam box containing dry ice to make them frozen completely.
You can also check a paper on JOVE (PMID: 4215994), which described freezing and cutting skeletal muscle tissues using this method in detail. But you may need to make some modifications based on the nature of the tissues you are studying.
Another option is fix the the tissues by formalin first then dehydrate tissues by highly concentrated sucrose before freezing tissues. In this method, you can embed tissues by OCT and freeze tissue using EtOH-dry ice bath. But you need to do antigen retrieval if you want to do IH or IF (different primary antibodies may need different optimal antigen retrieval methods) using those tissues. My friend in is another lab is handling tissues in this way but I don't know many technical details about it.
Hope it helps!
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Hello there,
The literature is plenty of examples of immunostaining in live cells, but is hard to find the same in live organotypic culture. Does anyone have experience with this? Is there (a priori) any caveat about this technique?
Thanks,
J
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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!
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I am trying to count the number of sperm cells in a fluorescent labelled image. After using the multi-point tool when I click measure the area column for all the selected cells is ZERO. I need that measure to calculate the CTCF(Corrected Total Cell Fluorescence) as
CTCF = Integrated Density - (Area of selected cell X Mean fluorescence of background readings)
I am neither using Thresholding (since I don't know why should I use it ) nor have I converted the image to binary. (Again. I don't know why should I use it )
Please Help.
P.S. Is this the correct way to quantify the fluorescent intensity?
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Multi-point tool marks the locations where you click. It's literally a 'location' (with sub-pixel precision), not a area. You may want to try the combination of regular ROI tools (like rectangle, oval, polygon, etc) and ROI manager (under Analyze->Tools->ROI manager). If you really want to select only one pixel for each cell, try wand (tracing) tools next to the multi-point tool. After you draw a ROI, simply press t to add it to ROI manager. BTW, I'm using Fiji/ImageJ.
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Hi All,
I am learning how to do immunostaining. Can you please share some established protocols for immunostaining breast cancer cells like MCF-7? How do we decide which primary and secondary antibodies to use? Once we decide the antibodies, how do we decide the volume?
Thank you in advance.
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Hi,
To do immunostaining for CELLS, we use immunocytochemistry(ICC) and/or immunoblot(western blot).
Regarding ICC, you can see this document https://images.novusbio.com/design/BR_ICCguide.pdf
Primary antibody selection is depends on your research objective, protein of interest.
Good luck