Questions related to Immunohistochemical Staining
I am using propidium iodide (Hi-media Product) but i want to know how much Concentration of propidium iodide is required to stain in 100 cells ?
In an article regarding IHC usen in myeloid sarcoma there was a report on a positive reaction to neutrophil esterase (Granulocytic sarcoma of the lips: report of an unusual case Badri Srinivasan. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2008;105:e34-e36), but i'm not sure if it is he same (synonym) to chloracetate esterase.
We are extracting total brain tissues and cutting sections via freezing stage microtome. Total brain tissues are fixed in 20 % paraformaldehyde + sucrose for 48 h and then transferred to %20 PBS sucrose. Until now, we were storing the slices at -20 degree in long term storage buffer. However, after staining some slices we got images with some kind of dirt and disruptions. The protocol for the long term storage buffer doesn't involve any anti-bacterial or anti-fungal but not all sections have this problem so we are suspecting that it is due to the storage time.
I would like to know what are the optimal conditions for storing the brain slices for long term or can we use another solution for long term storage of the sections? And how long can a brain tissue and a brain slice can be stored?
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.
In my imaging of mouse brains, there are fluorescent dots that appear in the image from confocal microscopy which are not any of the target cells in my experiment. I was wondering if using a secondary antibody from a donkey and a normal donkey serum in the blocking could potentially cause this?
I want to perform IHC for TFPI-2 in mouse tissue samples embedded in paraffin and although I find a lot of papers showing great stainings for human samples, I'm not finding it for mouse tissues. Somebody has tried to use a TFPI-2 with human reactivity for mouse tissue samples?
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!
I have been trying to do immunofluorescent labeling of IL-13 in mice brains (on animals injected with LPS to simulate inflammation) without much success. I am currently using 35 micron thick slices prepared by cryostat, the brains have been flushed with saline and fixed with 4% PFA. I've tried a lot of different techniques such as antigen retrieval, using TSA amplification, etc., as well as changed a lot of variables such as varying length of blocking, primary antibody incubation, etc. If anyone has had success staining for IL-13 or any other inflammatory cytokines such as IL-6 in mice brain, could you share what methods you used? Thank you!
I've been having issues with staining my postnatal day 7 rat hippocampal and lateral ventricle slices for iba1. They are 5µm thick, so quite thin. However, my postnatal day 21 rat slices are also 5µm thick and iba1 looks great on these slices.
According to the literature, there should be staining present in both of these areas so I'm wondering if it's worth either using a different secondary antibody or if the slices are just too small and thin at that age? I would love to hear any thoughts on what steps to take next.
Details about the slices for reference:
- Paraffin-embedded tissue sliced on microtome (5µm thick)
- 4% paraformaldehyde and PBS used for perfusions, post-fixed in 4% PFA for 24 hrs then soaked in 70% ethanol
- Coronal sections
- WAKO anti-rabbit iba1 primary AB
- Donkey anti-rabbit 555 (thermofisher) secondary AB
Thank you in advance,
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?
I would like to do IHC of HSV -1 TK in mouse tumors.
Can someone let me know where I can buy anti-mouse HSV -1 TK primary antibody for IHC?
Thanks a lot!
This is the result of immunohistochemical staining using paraffin-embedded mouse sections. Dewaxing, hydration, antigen activation (ph6.0 citrate, autoclave), blocking, and incubation of primary and secondary antibodies were experienced. DAB used for color development and counterstaining was also performed. The mounting medium was used with softmount.
The results showed artificial disturbances in the figure, like bubbles or crystals. Does anyone know what is causing this? What should be done to avoid it?
Good morning everyone (at least for me)!
Some questions for the resident ICC experts (and knowledgeable beginners!) out there- as my latest dabbles in immunocytochemistry have been disappointingly unfruitful. I have much more experience with immunohistochemistry, but unfortunately there are some snags I'm experiencing in translating my IHC experience to ICC.
1. When washing your chamberslides/coverslips do you apply the wash buffer "indirectly" onto the slide/coverslip itself (ie. using a chamberslide or coverslip and applying wash buffer to the corner of the dish/chamber to best not peel off cells) or do you immerse the whole slide/coverslip into a chamber/bowl/liquid-receptacle with the wash and then let it soak (akin to traditional IHC)? This can also sort-of apply to the initial fixation as well- do you dip the slides into a receptacle containing the fixative or do you apply the liquid directly onto the cells/chamber?
2. When using chamberslides do you keep the chambers on during staining or do you remove them before staining? If your slides don't have hydrophobic barriers then allowing all the slides to sit in the same antibody 'bath' could help with ensuring consistent staining- but it comes at the cost of losing the flexibility of being able to have multiple conditions (like no primary antibody negative controls) on the same slide. Lately i've been concerned that some of my chambers "leak" as I notice some chamber wells have less liquid in them following incubation than others- leading me to be paranoid there is not just leaking but contamination of one chamber to another.
3. What confluency do you typically wait for before progressing with ICC? I am currently doing an experiment on fibroblasts and I'm at a loss for what percentage I should let the cells grow for. I don't want them overly confluent, but I'm also concerned that if they're not 'confluent enough' they may not have good adherence to their slide.
4. How do you remove your liquid from your chamberslides? Do you turn the chamberslide upside-down and (gently!) shake the liquid out into a sink, or do you aspirate the liquid out every time? When working with secure tissue you can use all manner of roughness when immersing and shaking liquid off slides- but with cells I'm scared of them falling off due to their delicate nature.
5. Are there any common reagents used in IHC/ICC that you would *not* use for ICC? Triton is very commonly used in IHC/ICC but many places say that it can be too rough at times- could this roughness translate to 'scrubbing' cells off the slide?
I anxiously await the input that any professionals or beginners (like myself) may have and are willing to share. Advice, comments, tips, tricks, suggestions, criticisms, and thoughts of any kind are welcome and greatly appreciated! Likewise if anyone has any questions of their own I encourage them to share and contribute.
I promise to respond as soon as I am able to any response that comes in.
Thanks! Your help is immensely appreciated!!!
I have fixed some samples in 25x75 mm glass slides and I want to store them in a -80 C freezer. I put a layer of PBS on my glass slides and so I want the slides to stay flat in the -80 freezer. Any ideas on how I can store them? Will petri dishes be fine at such low temperatures? Please advise.
I am running an IHC experiment on mouse brain tissue that has been virally infected and I am looking to track infection by using and optimizing antibodies that detect immune cells, such as astrocytes and microglia etc.. In this experiment, I am blocking with normal goat serum (NGS) in PBS prior to using a goat anti-ALDH1L1 that reacts with mouse as a primary, and a donkey anti-goat as a secondary. This is the second time I run this with the same conditions and I dont see signal - it seems like the secondary antibody is not binding to/ picking up anything (similar to my negative control under the microscope). I am sure that there is antigen is the tissue - I previously used rabbit anti-GFAP and the signal of astrocytes is very strong and visible, so I would expect very similar outcomes with anti-ALDH1L1. Could this be due to the use of a blocking serum from the same species and the primary antibody and not the secondary?
I am currently re-running an IHC with formalin fixed paraffin embedded sections of mouse brain sections. I am using a primary anti-Iba1 IgG1 antibody that is raised in mouse and a secondary goat anti-mouse IgG antibody. My first run with these antibodies showed a high degree of background staining with low specific signal - I know what specific signal looks like with these sections since I had tried a different anti-Iba1 antibody in the past on these sections and I know that there is an inflammatory response. Would this have anything to do with the fact that my primary is IgG1 while the secondary is IgG - i.e., picking up what's not supposed to be picked up?
We perfuse our mice with PBS followed by 4% PFA, harvest brains and allow to fix overnight (20 hours) in 4% PFA before moving brains directly into 30% sucrose for 24-48 hours until tissue has sunk. Then the brains are frozen on dry ice into OCT and sectioned in a cryostat. The 20um sections are kept long term via free floating in a 24 well place immersed in our cryoprotectant recipe below:
STEP 1 Prepare PB (0.1 M phosphate buffer pH 7.2)
1. Prepare sodium phosphate dibasic stock (0.5 M Na2HPO4) by dissolving 35.5 g of sodium phosphate dibasic in a final volume of 500 mL of H2O.
Some crystallization will occur when the solution is stored at 4ºC. Warm on a hot plate and stir until the crystals dissolve.
2. Prepare sodium phosphate monobasic stock (0.5 M NaH2PO4) by dissolving 30 g of anhydrous sodium phosphate monobasic in a final volume of 500 mL of H2O. For NaH2PO4.H20, measure 34.5g.
3. Prepare 0.1 M sodium phosphate dibasic: Put 80 mL of sodium phosphate dibasic stock (0.5 M) from Step 1 in a beaker and add H2O to give a final volume of 400 mL.
4. Prepare 0.1 M sodium phosphate monobasic: Put 30 mL of sodium phosphate monobasic stock (0.5 M) from Step 2 in a beaker and add H2O to give a final volume of 150 mL.
5. Bring the 0.1 M sodium phosphate dibasic solution from Step 3 to pH 7.2 by adding as much as needed of the 0.1 M sodium phosphate monobasic solution from Step 4.
The resulting solution is 0.1 M phosphate buffer pH 7.2.
STEP 2 prepare Cryoprotectant (Use for Immunofluorescence)
To make 1000 ml cryoprotectant
• Sucrose ----------------------------- 300 g
• Polyvinyl-pyrrolidone (PVP-40) --- 10 g
• 0.1M PB ----------------------------- 500 ml
• Ethylene glycol --------------------- 300 ml
Add PVP-40 to 0.1M PB. Stir to dissolve. Slowly add the sucrose to dissolve, and then add the ethylene glycol and bring the final volume to 1000 ml with 0.1M PB. Store at -20°C.
I am looking to know how this cryoprotectant can affect tissue overtime for lipid based targets, specifically MBP? For our longer kept tissue ( > 1 year) we are having an unsucessful time staining for MBP via IF and the black gold II kit. I have suspected the cryoprotectant for a long time, seemingly the longer kept tissue doesn't stain as well; When i compared floating sections kept in cryoprotectant vs directly adhered sections, dried and then stained, the color outcomes for the black gold kit were very different. Directly adhered had more dark purple/blue/red while the tissue that had been exposed to cryoprotectant was more black/grey (diverging from expected color results but still identified tracks appropriately).
We have tissue that can't be recollected and i need to figure out how i can save the tissue currently in cryoprotectant for myelin staining. Neither IF myelin nor black gold II kit staining is working on this older tissue, which we should be seeing ample amounts of ( by WB analysis of mice collected from this group, myelin is ample) but staining will work on more newly collected tissue exposed to cryoprotectant.
Does anyone have any suggestion or publication about the effects of this type of cryoprotectant on targets post fixation? I was wondering if the direct immersion in 30% sucrose was an issue but again, this appears to be an "over time" problem for our sections.
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?
The attached images are at 40x total magnification. (Please excuse the lack of a scale bar; it disappears during file conversion.)
Blocking for endogenous peroxidases does not solve the issue. (To check this, I included a 30 minute incubation in 3% H2O2 before antigen retrieval.)
We use the at-least 98% p.a. 3,3'-Diaminobenzidin Tetrahydrochlorid powder from Carl Roth (Order # CN75.2), and the buffer for our DAB solution is 0.175M sodium acetate with 5% imidazole.
Any suggestions for how to solve this problem would be very much appreciated.
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. ?
I performed multiple c-Fos IF on DVM RVLM sections after seeing no reaction on the previous ones and to test if there is a problem with our antibody or is there any effect of temperature on the tissues. Surely +4 should be preferred for incubation but I am wondering if I also need to use shaker at +4 (if I can manage to do that) to get better results and no damage on the tissues.
I hope that you are all doing well. My lab is looking to stain NMDAR1 using immunofluorescence (free-floating tissue - PFA fixed) in CD1 mice. However, we are having difficulties as we cannot find an antibody anywhere! I was wondering if anyone has used an NMDAR1 antibody that they could recommend (host doesn't matter - most are rabbit anyways).
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:
iNOS --> Alexa Flour 488
Iba1 --> Alexa Flour 594
Arginase 1 --> AF488
Iba1 --> AF 594
I couldn't find a resource on how to quantify this, any help would be appreciated.
Which primary antibody is better to IHC staining collagen III and collagen I in mouse skin? I need Cat Number.
Hi all, I am performing IHC to see the HMGB1 distribution in my C57BL/6 mouse tumor tissue. I followed the standard protocol from Abcam for deparaffinization, rehydration, and staining. But I get completely no staining on HMGB1 but only nucleus counterstain by hematoxylin (As figure shows). The primary antibody is from cell signaling and works for IHC, and the secondary antibody is HRP-conjugated which targets the host of the primary antibody. There are few concerns I have about my IHC procedure.
For antigen retrieval steps, I initially used the microwave to boil pH6 citrate buffer but the buffer rapidly evaporated and left dry slides in the microwave for over 10 minutes. Then I changed to use a 95-100 celsius water bath to heat the citrate buffer with slides. I'm wondering if the antigen was destroyed in these steps?
I also checked my DAB substrate kit, I found that the diluted DAB substrate followed by protocol showed transparency with very few brown colors. I'm wondering if I should allow the reaction between DAB and HRP-antibody longer than 10 minutes?
It is my first time doing IHC. Could someone please provide some suggestions on why there is no DAB staining on my slide? Thank you very much.
I was just wondering whether it is possible to make Cresyl Violet/Nissl (or a similar stain to determine location of implanted electrode) in the lab instead of buying it?
I have a colleague that prepares their own Golgi solution, so I wondered whether the same could be done for Cresyl Violet.
EDIT I found some Methylene Blue in the lab. Could this be used to stain tissue in a similar way? Or are there any other lab alternatives?
I would like to know any simple stain or test bu which i could stain or quantify only the EPS or biofilm present and not bacteria present in it.
I have a question about rabbit GC B cell. I am working with respect to virus and how it affects rabbit GC B cells. However, i am unable to get any good antibody for GC B cells. Tried BCL6 but the clone i selected is not staining well. I was hoping to get some answers here.
Thanks in advance.
I have brain tissues that are fixated for a few years and I plan on staining them using fluorescent Neu-N antibody. I know that it usually does not work without antigen retrieval. I could not find a reliable protocol for that. What do you think is the best way of working with those tissues? A detailed protocol would be much appreciated.
Hello everyone, I just started learning IHC staining and I have some frozen sections to practice with in a 96-well plate. I'm having trouble mounting the sections on the slide, a colleague told me to use just 1xPBS to float the section (she uses PBS + triton, but told me I shouldn't begin with that) and then mount it using a brush on the slide, but it's much harder than it looks, and the sections always sink and I screwed many sections because they are so delicate. Do I need to change the solution? I watched many videos for the technique, but I don't know if there's a better solution, or any tips generally.
I have dissected 2-3cm intestinal segments, flattened and stretched on a platform, fixed in 4% PFA for 24 hours, and dehydrated in 0.03% sodium azide for a maximum of 4 days (is this too long?). After this, what are the best solution and temperature conditions to store this tissue if needed for immunohistochemistry?
Also - what is the best method by which the myenteric plexus and submucosal layers of the gut can be isolated from these whole mounts for histological staining? Thank you!
I would like to stain a tissue for different markers, and as I there are not enough colours that I can use, I would like to do it sequentially. Thank you in advance!
I was wondering if anyone knows an antibody that can be used for staining microglia in zebrafish? I am familiar with 4c4 and L-plastin antibodies, but these are not perfect for me. Is anyone familair with, for example, a P2Y12 ab that works in zebrafish ?
This is an IHC staining for CD142 figure for perivascular adipose tissue. I wonder what are these cells containing particles among mature adipocytes?
Will they be mesenchymal stem cells or immune cells resident in adipose tissues? And what are those particles within these cells?
I've stained some bright field Arginase-1 Hematoxylin slides, but I'm having a hard time finding any computed histology predicates for evaluating the intensity of the cytoplasm verses the nuclei; as everything seems to be done manually.
I've attached some 20x images of the slides. But I'm concerned about edge detection of the cell and haven't had much luck with segmentation in ImageJ. I lose a lot of edges in-between some of the cells.
- Does anyone have any other software or ImageJ plugins you'd recommend?
- Any specific computed histology guides or methods of hepatocyte specific antigens or something similar?
- Any other guidance?
Thank you for your responses.
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
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)
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,
I am using the clone from Biolegend but is experiencing difficulty in optimising the antibody. I have tried using various dilutions as well as various antigen retrieval methods but to no avail.
Does anyone have an IHC protocol for detection of a his-tagged protein in mouse tumor tissue (cryo, not paraffin)?
I'm trying to stain for a protein that contains a 6His tag and accumulates in tumor tissue (human tumor xenograft grown in mice). I used the M.O.M. blocking kit from Vector Labs to block endogenous mouse IgG, then incubated with the mouse penta-his primary antibody. The secondary antibody used was the Vector Labs Impress HRP-anti-mouse antibody. The problem is I get really high background signal, even in the negative control (no primary antibody). It seems the only species of penta his antibody is mouse derived, so maybe someone knows a source of rabbit, goat, or donkey penta his antibody?
I'm looking to do some IHC/IF staining of X. laevis brains that have been frozen in OCT media for cryosectioning. I understand how to do the cryosectioning, but I am hoping for help/protocols that will assist with the following steps:
- Preparation of Chrom Alum Gelatin-coated slides
(I've had issues with my tissues adhering to the slides before, so my PI suggested this step)
- Antigen retrieval
(Is this step necessary? I'm working with brains that have been fixed in MEMFA (formaldehyde), so I'm not sure if the protocol will require this or how to do it.)
- Immunofluorescence staining of sections once they're on the slides
(Could someone walk me through the best way to add your antibody solutions to the slides without damaging the tissues? Do you prefer a bath that you dip slides into, or do you use a hydrophobic pen to create a barrier and keep the slides in a humidified chamber?)
- Using parafilm to create a barrier instead of a hydrophobic pen
(I've heard of this before, but I'm not sure exactly how to do it, so I would love any tips/visuals/advice anyone might have!)
Thanks so much!
Any ideas on how to quantify a non-nuclear immunofluorescent signal in adipocytes? I had the idea to do a watershed on inverted image to "locate" the center of the empty space where lipid droplets used to be, and then maybe estimation would be possible by calculating the AUC of the "first peak" for the plot profiles extending form the "point-maximum" outwards. In case it's not clear, I'll attach an example of what I had in mind with a schematic and what the output would look like in a single inverted HE adipocyte (the same should work for IF). Is there a package in ImageJ/Fiji that could help with this or maybe another way this could be automatized? Somebody working with adipocytes might know? I know there are some packages and even stand-alone stuff for adipocyte morphometry (AdipoCount?) sop for those working with these, do you know if it could be done in any of the software available or has to be custom-made?
Also, this is what first comes to my mind, but there are probably better ways of estimating the cytoplasmic/membrane expression as this provides the output based on rather small sample of the membrane (6 profiles) with each sample suffering from bias based on expression signal from surrounding adipocytes, although I guess this is not extremely important as you are usually interested in them as a group rather than in the expression in single cells. But if you have other ideas I'd be really interested..
Some protocols measure the optical density of the immuno-reactive cells while others use scoring system. What is the best method?
I'm planning on doing oligodendrocyte staining on frozen and paraffin brain sections of postnatal (up to 2 weeks old) and adult mice.
I was wondering which antibody would you recommend for cytoplasmic staining of oligodendrocytes. Unfortunately, my other marker is nuclear, and would therefore preferer a cytoplasmic or membrane marker.
I know that there are a few commercial antibodies, however, I can't deicide which one would give me the best signal to clearly distinguish oligodendrocytes from other cell types in the CNS.
Thank you for your suggestions,
All the best
I am working with some colleagues on historic FFPE sections from multiple sites that have been (unknowingly) cut onto different types of glass slide.
We have used Thermo SuperFrost Plus (J1800AMNZ) and Thermo Polysine coated (J2800AMNZ). Both are positively charged.
I know that positively charged and non-charged slides may display different staining properties; I wonder if anyone has any experience with different charged coatings and whether this is something we need to be mindful of regarding quantitative analysis?
I wish to a stain a HA-Tag protein by IHC from cell-lines implanted subcutaneously in mice. The protein is very unstable by nature and expressed at low levels. Is there any good well-tested anti-HA tag antibody for IHC of such proteins?
Any recommendation and direction to specific protocol is highly appreciated. Thanks.
I want to state that I have limited training on staining with alexa fluor 488.
I currently have to detect CD31 for angiogenesis on mice sectioned tissues. And I bought alexa fluor 488 anti mouse CD31 from Biolegend.
For the most part I know I need to block the tissues (5%BSA), and then incubate tissues with alexa fluor 488, followed by Dapi incubation.
But I don't know what concentration to start with. The alexa comes as .5mg/ml. I normally used 20uL in 1mL for in situ zymography staining.
Also, if anyone could recommend how to quantify the amount of protein/fluoresence for my images, that would be great.
Thank you for taking your time to answer,
I've been doing IHC for years on tissues on slides and this is my first experience with staining free floating brains via IHC. Both primary antibodies (diluted 1:2000 - 1:5000) came up completely brown in 30 seconds of DAB. Can anyone suggest reasons why this may have occurred?
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!
I have 3 groups of animal apart from PBS controls where i treat them under different immunosuppressive conditions. when the animals are subjected to partial or no immunosuppressive conditions, i do see the nuclear expression of BCL-6. When the animal is completely immunosuppressed, i do not see nuclear expression of BCL-6 rather than cytoplasmic. Also, the staining is very faint, eventhough i tried different concentrations. Can i know what would be the reason for cytoplasmic exprression and why do i see only faint staining even with antigen retrival?
I have attached some images for reference.
thanks in advance.
In IHC for double staining i use DAB and AP. DAB works fine but AP after mounting is a headache as i see precipitates and also sometimes dissolves.I have tried drying the slides a bit and then mounting with the aqueous mounting media. I first wash it with water or rather rinse it with water after NBT-BCIP step and then mount.
How do i overcome this.
Thanks in advance.
Does eosin stains collagen in IHCs in typical H&E staining? Please share your thoughts below as comments. Please add a reference if possible (much appreciated).
I am currently storing my fixed slides at 4 C. These slides tend to dry out. Are these slides still good to use? can these slides be revived by adding liquid? I have attached some images .
I am working with rabbit spleen and as it is difficult to obtain antibodies against rabbits as most of them are raised in rabbits. I had to purchase a few antibodies raised in rabbits. But as expected, i am getting background staining in IHC-P. Is there any way that i can reduce this background staining.
PS - Little help with rabbit specific B cell markers are much appreciated.
Thanks in advance.
I'm analyzing some TMA IHC cores using an scoring method that combines intensity and proportion of stained cells. I would like to know if is there any protocol to quantify percentage of DAB stained cells (the markers I'm using stains both cytoplasm and nucleus) and if is there any that can be automated (using ImageJ maybe?) since I have plenty cores?
I found this article
Hello, we are developing a protocol for combining FISH and IHC on Drosophila embryonic brain, using FISH to localize mRNA and IHC (2 antibodies) to mark neuropil and neural tracks. We thought about mixing anti-DIG antibodies with one of the IHC antibodies, but our usual protocol for IHC calls for incubation in antibody for one to two days at 4ºC, and the protocol I found for anti-DIG says to incubate for at least 4 hours at RT. So now we're planning to separate the two processes, but we don't know which one we should do first. Any protocol or suggestions will be appreciated!
If it helps, here are some of the materials we're using:
Anti-DIG-Fluorescein Fab Fragments from Roche
IHC blocking: 100ul normal goat serum + 900ul PBT
IHC protocol: PBT wash (2x20min) -> block (30min) -> 1º staining (O/N) -> PBT wash (3x20min) -> block (30min) -> 2º staining (O/N) -> PBT wash (30min)
i am having difficulties with staining for Phospho TAU in KA-induced animal model.
I am using free floating immuno histochemistry method.
I am using AT-270, At-180, AT-100, AT-08 antibody (1:200, 1:500, 1:1000)
I have tried antigen retrival (sod citrate ph6)
i have tried 3%H2O2
I am trying blocking next lets see..
And Doing DAB staining.
Can any one help me ..or is there any specific technique????
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?
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).
I am planning on performing expansion microscopy on relatively thick (300 um) tissue slices and doing a post-expansion stain with some nanoparticles. It seems that the expansion process should make the tissue more permeable, but I am having trouble finding a direct description in the literature which supports or goes against this intuition. Does anyone know whether or not a post-expanded tissue has greater permeability than an untreated fixed tissue sample? Note: my nanoparticles are rod shaped and have dimensions of 27x60 nm, but I could also try spherical 5 nm nanoparticles if needed.
I've been trying various iba1 antibodies with either Alexa 488 or 647 and am getting a lot of background and have never once seen anything close to resembling a microglia. I've tried blocking with various combinations of 0, .3 or 3.0% milk, 0, .3 or 1.0% BSA, with 4% normal donkey serum and nothing has worked.
Any suggestions would be very much appreciated. Perhaps there are better membrane-bound proteins I could stain for?
Hello! I am a student and I am trying to do immunohistochemistry staining. I am wondering if there are specific markers for cranial neural crest cells only. I found many for neural crest cells in general, but am unsure if there is any which is specific for CNCs. Thank you!
I'm taking over a project where all the primary ab have already been added to the slides. The person before me didn't document his work very well, I know he used donkey serum before adding the primary ab and then some of them he used goat serum. No idea which ones.
So I'm wondering if it makes a big difference if I used a anti-mouse IgG (secondary Ab) raised in goat if he incubated the primary ab with donkey serum since the rule of thumb is to use the same species as ur secondary ab...I can't find any literature on using serum and ab of different species. (primary ab was raised in mouse and it's on a mouse tissue).
Currently, we have been performing angiogenesis assays on 15-well Angiogenesis µslides (Ibidi) and are attempting to find a way to perform immunohistochemistry on the resulting cells. Has anyone successfully been able to remove the cells from the wells in order to embed them and section them?
Thank you in advance.
Paraffin embedded staining is easy, but someone 'mentioned' that they thought it was harder in frozen cryostat sections. I would like to collect for frozen to expand the later uses of my tissue. Could anyone please provide a protocol or guidance on CC3 in frozen (15uM cryostat cut) section. Thanks!
I would like to collect the cervical spinal cord sample from rat following retrograde tracer injection in the forelimb muscles. After collecting the sample, I would like to find out the corticospinal tract changes. Can anyone please help me to find out a protocol for IHC and 3D-Z.1light-sheet microscopical view of the cervical spinal cord.
Thanks in advance.
I have flash frozen some brain slices in OCT with isopentane, cooled by liquid nitrogen
I have mounted some on slides using a cyrostat however I am finding that the tissue dissociates from the slides very rapidly.
I just wondered if anyone had any advice? It was deemed unsuitable to fix the tissue with PFA before storing in the -80
In case that, it is difficult to get animal serum from commercial vendors. Have you ever tried to collect animal serum to be used for blocking in immunohistochemistry ? If so do we have to sterilize it before use? and by which method?
Thanks in advance