Immunohistochemical Staining - Science method

Can anyone suggest me that how much number of cells are required for flow cytometry and what is stock concentration and incubation time of Propidium Iodide?

Hello Moses Grimaldo Carjevschi

Histologically, granulocytic sarcoma (GS) undergoes various morphological changes in the cells. It is composed of immature cells of the neutrophil granulocytic series. The infiltration of immature, poorly differentiated cells with round to oval nuclei is the predominant finding. Crystalline, rod-like, intracytoplasmic acidophilic bodies called Auer rods are sometimes seen. With these histological findings, it is hard to distinguish GS from other lymphomas including histiocytic lymphoma, lymphoblastic lymphoma, Ewing's sarcoma, lymphocytic leukemia, undifferentiated carcinomas, and primitive neuroectodermal tumors.

You may refer to the article attached below.

Anti-lysozyme and chloroacetate esterase can be used to diagnose GS.

There are two types of esterases, specific and non-specific. The specific esterase will enzymatically hydrolyze naphthol AS-D chloroacetate liberating a free naphthol compound. This then couples with a diazonium compound, forming highly colored deposits at sites of enzyme activity. This enzyme is usually considered specific for cells of granulocytic lineage. This specific esterase stain, chloroacetate esterase (also called Leder stain), stains neutrophils, neutrophil precursors, eosinophils, and mast cells and confirms the granulocytic nature of the tumor.

On the other hand, the nonspecific esterase activity (alpha-napthyl acetate esterase) is seen in monocytes. The cells of granulocytic series are negative for the nonspecific esterase activity.

So, in the article regarding IHC used in myeloid sarcoma in which there was a report on a positive reaction to neutrophil esterase means that the author is referring to the specific esterase namely, chloroacetate esterase.

Best.

Dear Alexander Trofimov , thank you for the information you provided.

I was wondering if it is crucial to replace the storage solution periodically?

Please check abcam company it might be useful for your experiment

Positive control

Hello Deven Gopinath

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

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

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

Best.

I think I could recommend TFPI2 Antibody (CSB-PA023438DSR2HU) to you. Here is the page:

https://www.cusabio.com/Polyclonal-Antibody/TFPI2-Antibody-11080650.html This antibody had been applied in WB ( mouse samples), IHC (human samples) successfully.

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

Thanks Irena,

Do you think or have you experienced labelling soluble interleukins, that they diffuse in the intercellular space i.e. extracellular matrix? Or do you usually find them mainly within cells?

Hey Melissa,

You got signal in one age and none in the other within the same staining batch? That's really weird.

I don't think that the thickness is a problem. I'm staining even thinner sections - 3 microns - (mouse brain) with Iba1 and get nice signal.

If it's not biology (sorry, I have no clue about 7 day old rat brain), you could try to tweak your protocol.

You could include TritonX (0.1 - 1%) to your blocking or staining buffer (but beware, it then easily spills over the Pap/Pen). Or, what I suspect is more likely to help, perform an antigen retrieval step before the primary antibody. E.g. by boiling in citrate buffer (we're using pH 6 for 20 min boiling time in the microwave).

Best wishes,

Theresa

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.

Did you happen to have any luck finding a good antibody?

before mounting sections you should wash your lam with passaging through very strong base like KOH and then put them in D-water and next strong acid like HCL and again DW in this way your lam will became very clean then instantly put them in fresh filtered gelatin solution and leave them dry for a night and make sure not became polluted by covering them.

after sectioning and dewaxing, passage sections in another bath and make sure sections perfectly dewaxed when mount on lam also surface of lam most not encounter with wax specially under of mounted section.

use sampler for Entellan dropping on lam and make sure on bubble remain under lamella by pushing lamella.

do not heat Entellan or gelatin coated lam because distribution became disrupted and making bubble.

good luck.

Block deparaffinized sections with blocking buffer. Than Use antigen removal by the method you like, there are several methods. Antibody company may suggest you, read the sheet comes with primary ab.

Composition of blocking buffer:

o Blocking buffer (BB) Preparation (PBS-T containing 2%BSA and 2% animal species serum (serum always from animal species in which 2˚ Ab is raised)

Mix 25ml PBS-T+0.5ml animal species serum +0.5gm BSA and leave at RT to dissolve

o Preparation of PBS-T solution (PBS containing 0.05% Tween-20):

To a bottle of 500ml PBS placed at RT, with 1ml tip, add 250ul Tween-20(Final concentration: 0.05%) (cut the tip to avoid bubble). Mix by shaking bottle 4 times up and down and leave the bottle at 30 min at RT

After 10 minutes of blocking section, you use primary ab,

wait for 30 min to 1 hr, (dilute primary ab in blocking buffer)

Wash antibody with blocking buffer 5 min 3 times

Than add secondary ab (dilute secondary ab in blocking buffer)

Note: In blocking buffer, serum should be from species in which secondary ab is raised

5. It depends of the cell type. I tested Triton on stem cells and it washed them out. All my antibodies worked well with just normal serum (no Triton) on SC and also neuronal cell lines.

It is better to use to use polystyrene petri -dishes at -80 C

Hello Karim Zedan

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

In your case, you should avoid using goat serum for blocking if you are using a goat primary antibody with anti-goat secondary antibody.

Use serum that is from the same species as your secondary antibody. You could also use BSA for blocking.

Best.

You are right. The high background could be due to nospecific bindung of anti-mouse sec Ab. If possible, use primary Ab dierectly tagged with marker (flourscence or HRP) i.e. one step detection method of IHC.

من خلالها نقل الدهون والفيتامينات الذوابة في الدهن بعد امتصاصها على شكل مستحلبات دهنية هي الدقائق الكيلوسية

ربما يكون ذلك

Hi Amanda, I would strongly recommend this rabbit anti-Olig2 antibody from GeneTex (REF: GTX132732). I have used it to perform double BrdU/Olig2 staining in rat V-SVZ!

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.

Thank you, Gail Stonebarger! Those three rinses are part of our protocol also (we do 5 minutes per rinse). I actually did find that tablets work better than the powder! They are just sadly too expensive for the amount of solution our lab regularly uses.

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.

I've known some people to use a shaker, but I had results with just leaving the samples in the +4C, my first thought would be to check the antibodies whether they are old or perhaps were kept improperly. It wouldn't hurt to try with a shaker but not sure how much it'll help

Hello Kevin,

CUSABIO provides the custom of NMDAR1 antibody. 200ug * antigen (positive control) and 1ml * Pre-immune serum (negative control) are offered.

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

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

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

Hi, arigo's ARG20786 Collagen III antibody has been validated by paraffin-embedded sections from Col3a1 mutant mice. Please refer to the datasheet at https://www.arigobio.com/anti-Collagen-III-antibody-pre-adsorbed-ARG20786.html, and the reference from doi: 10.1101/2021.06.12.448202.

The center of photomicrograph appears to contain mostly necrosis which probably resulted from dead cells or incomplete fixation of tissue. Moreover, staining of nucleus by hematoxylin should be blue and not red as is here.

I suggest IHC staining of the tissue section with antibody to pan keratins, provided the tumor derives from epithelial cells, may reveal any problem with fixation with 10% buffered formalin or 4% paraformaldehyde of tissue.

Was the antibody to HMGB1 you used recommended by the vendor for FFPE tissue section.

In order to avoid drying up of sample on slides, one could place (1) the slides in a plastic staining rack in the antigen retrieval solution, (2) the rack in a plastic pressure cooker, and (3) the pressure cooker in a microwave oven for the purpose of AR.

Sounds 2.5N HCL is very concentrated itself for tissue IHC to begin with. You may tell whole protocol in detail than someone can answer you.

I wonder too if there meanwhile has been any progress with regard to the initial question matter...it would not only be of benefit for those who perhaps were seeking a / looking for an answer to this post, I unintentionally came across after long time passed - only due to an other request to my private e-mail-address.

But most fortunately I wish that the problem has been solved meanwhile to the satisfaction of Kristie Leigh Smith .

FilmTracer SYPRO Ruby biofilm matrix stain

Thank you very much Lora Benoit . I tried PNA from vector labs and unfortunately its staining for T cells and am also getting non-specific staining.

Any other GC markers reactive with rabbits would be helpful.

Thank you in advance.

Naren

Brain tissues even when they are fixed in 10% NBF for 24 hours are less robust than those derive from epithelial origin, for example. However, brain tissue section of 4 micron of thickness placed on Superfrost Plus glass slides, air-dried preferably overnight and baked at 50 to 55 degree Celsius, depending upon the characteristics of the antibody being used, for 3 to 4 hours, could be easily immunostained on any of the commonly used commercial IHC platforms.

Good luck.

Finally it is a kind of expierence. The more you mount the better will be the result.

Some tipps: use gelatine coated or Superfrost plus slides as already mentioned. Instead of PBS/TBS I recommand a 0,45% NaCl-sorution with a drop of liquid soap (or Triton) in a petri dish. I prefer NaCl solutions because the salt cristals are easierly soluble then the buffer cristals after air drying.

If you are using sections thinner than 10µm , cut the tissue on a Cryostate and transfer the sections directly on slides.

Check https://www.niehs.nih.gov/research/atniehs/labs/assets/docs/q_z/revised_guides_for_organ_sampling_and_trimming_in_rats_and_mice_508.pdf

Hi janmaris@Janmaris Marin

you showed that you got good results of destaining the first antibody, I wonder if the striping buffer that you used was glycine-SDS（pH 2.0）in the Daniel Pirici article which your mentioned above.

Best wishes,

Lixin

IBA1 from Wako-Chem Cat: 019-19741 is reported specific for zebrafish

well these are adipocytes for sure !

You might find CellProfiler useful. Agreed, the cell borders are very faint. If you exclude nuclei and quantify staining within cells, might not need to segment individual cells. Good luck.

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

Xingzhou Wang Unfortunately we reached a dead end and for other reasons, we could not continue on this track of the project.

I am curious which goat polyclonal anti-6his antibody did you end up in using from abcam? Thanks!

1- You can use positively charged slides for better attachment. Or you can coat your slides with agents like gelatin or lysin. Both protocols are quite easy to find online and easy to oparate. Also make sure your slides are completely dried before you start.

2- Yes you should perform antigen retrieval since you use a formaldehyde based fixation. For that, you can use enzymes or you can boil your sections in buffer solutions like citrate buffer or tris-EDTA buffer.

I mostly prefer using citrate buffer pH:6.0. Prepare the buffer, put your slides in a challet filled with buffer solution and microvawe for 5 mins (for three times). Let your slides cool down a bit between microvawe sessions.

But if you have some problems with adhesion, you may prefer trypsin for retrieval. You can find the protocol for that in the link: http://www.ihcworld.com/_protocols/epitope_retrieval/trypsin.htm#:~:text=The%20trypsin%20based%20solution%20is,enhancing%20staining%20intensity%20of%20antibodies.

Note: Since you're working on frozen sections skip first two steps of the protocol. The antigen retrieval step is the 4th step alone.

3- Hydrophobic pen+humidity chamber works fine for me. Mark your sections, drop antibody solutions, place a plastic coverslip onto the section, make sure there's no bubbles on the slide and incubate them in humidity chamber at +4C overnight. Don't worry, if you place coverslip w/o bubbles your sections wont dry out. I use very small amounts of antibody solutions (like 20-30 microliters) and it's working without any problems.

4- I believe if you use parafilm instead of h.phobic pen, the parafilm may melt down in antigen retrieval step. If you don't have a h.phobic pen, you can use a diamond pen or some sharp equipment to scratch the surroundings of the section.

Hi Jan,

Check this method out:

Whole-slide image analysis outperforms micrograph acquisition for adipocyte size quantification. Adipocyte (2020), 9:1, 567-575.

There are a few more methods along those lines. Good luck.

@Sathya Srinivasan

Dmitry Zinovkin I want to measure the expression of some proteins by cells

Dear Fran,

It depends on the differentiation stage of the cells:

If you want to stain OPCs, you could stain with an antibody against PDGFaR. For oligodendrocytes, MOG, Opalin or CNPase works, mature oligodendrocytes can be detected with MBP.

I would not recomment antibodies against glycolipids, such as GalC, O4 or O1, for frozen tissure. Olig1 and Olig2 are transcription factor, therefore you don't want to use them.

Good luck,

Sebastian

Good day. Actually , positive charged and polylisine coated slides are used for immunohistochemistry because it prevents fall of section from slides during rinse in buffers heat antigen retrieval. It doesn't play role in quantitative analysis of results

Many companies have Tag antibodies, and the quality is very good. HA Tags are commonly used antibodies. There must be what you need. You can consult an antibody company.

Which ABCAM anti-CD31 do you use for mouse tissues?

Hi guys, I was following this chat. COuld you recommend a protocol for frozen sections (human brain tissue) for free-floating immunofluorescence? I did only free-floating immunofluorescence for mouse tissue. The tissue I have was fixed in PFA4% and stored at -80°C. Normally we do cryosections and on-slide IF.

thanks in advance for your input!

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.

Thank you very much dani. It was very helpful. Any thoughts on why the staining would be cytoplasmic and not nuclear as it should be?

Renee R Donahue once i develop with the chromagen NBT-BCIP, i rinse with water and take out the excess water and mount with aqueous mounting medium that i bought from sigma.

sometimes i get these precipitates or sometimes the developed stain dissolves.

any tips?

thanks again

Thank you..!!

here are some reliable suggestions, my team has been working on these methods for 10+ years and i used these methods since 1997:

1st, some of the weird-looking areas on your 3rd image are salt crystals or droplets of moisture, they will be gone when you dip your slide inn PBS or water, no need to worry.

2nd, if you have paraffin-embedded sections of PFA-fixed tissue, they are safe at room temperature in your slide box for years. Some people may store them at-20 but the antigens immersed in paraffin and also fixed with PFA are stable for years.

3rd, looks like you have thin section of frozen tissue, lightly fixed with PFA, it is rather fragile and crumbles a bit. Use 10-14 micron sections for frozen sections.

when a section comes off the blade, mount it on a slide, put slide on a tray at room temperature, continue cutting until you're done. By that time they are usually dry. Put them in a slide box and store at -80. Do not do repeated thawing-freezing, when you need to select a slide from a box, pull out a box from -80, put it on a tray with dry ice (if you need more time) and select needed slides. Otherwise, quickly open -80, keep slide box in -80 on a cold shelf, quickly select needed slides while keeping the slides at below zero C (on a -80C shelf), , put the box back in -80. some people store frozen slides for some time (1week -1mo) at -20, i do not recommend long term storage at -20, but short term - no problem.

So, long answer to how you store -air dry them as above -and stick in -80C. It's ok if you cut and leave slides to dry overnight, then collect next morning and put in -80. If you're doing in situ, you have to be more careful how you store the slides. For shipping, experienced people ask to vacuum dry your frozen sides (paraffin slides are ok as is) using a regular lab vacuum desiccator. An alternative way of storing frozen sections, which are not falling apart (and this is typical for those working on brain or spinal cord) is to collect sections in 48-well or 24--well or 6-well etc. dishes in the mix of ethylene glycol, 10x PBS diluted to 1x, and glycerol (i.e., you keep them in solution, not mount on slides), several consecutive sections can be in the same well, or, if needed, each section can be in a well of 96-well plate, this works). Use a paint brush to move sections from a blade to solution. Then keep trays of slides (96-well, 48-well, 6-well etc) at -20 to -25, the cryoprotective solution wont freeze and will keep sections cold, and when you pull them out, you rinse in PBS and then stain so called floating sections and after finishing up 1 and 2 antibody, mount on a slide. You can store sections like this for 1 -2 years and longer. Never tried for 5-10 years :) Hope it helps. Good question by the way For references: https://www.ncbi.nlm.nih.gov/myncbi/1l14BdjIrshQ9/bibliography/public/

I guess the cleanest way would be to directly conjugate your detection moiety (e.g. HRP) to your primary antibody. Any detection method that relies on detecting your rabbit antibodies by a secondary antibody recognizing rabbit immunoglobulins will also detect endogenous antibodies in rabbit spleen.

Update: With all the trails and fails: iam still having difficulties with the staining of Tau in my Rats.

May be the chemical iam trying is not inducing tau hyperphosphorylation.

I need to think of new ways until iam 100% sure of it.

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

I have been using expansion microscopy for 2 years now and in my personal experience, small things such as DNA oligonucleotides can be introduces post-expansion (typical length scale of 13 nm). However, longer incubation and wash steps are needed to hybridize oligo's and remove non-bound oligo's. Knowing this, I think your 5nm particles can penetrate for sure to the center of the expanded sample, however you might need to incubate longer to cross the larger distance. However, the exact pore size of the expanded polymer has not been described and also depends on the amount of crosslinker added (for a typical expansion gel this is 0;.15% N,N methylenebisacrylamide)

Hope this helps.

try vector trueview, autofluorescence quenching kit with DAPI (Cat# SP-8500)

Hi Noelle, did you find a good marker for cranial neural crest cells? I am interested in this.

Our laboratory has (inadvertently) done exactly what you describe - that is, block in donkey and use goat secondaries. We also were not able to tell a difference between this and a round of staining that used the same species for both. I don't think there is much to worry about, but for sanity purposes it may be nice to use the same species moving forward.

Take a look at my publication on this topic. I developed a technology especially to do this. I created a well insert that can be cryosectionned. Search on the internet, 3D Cryo well insert.

Dear Boobi,

Please, could you send me the protocol in case you did it.

Thanks in advance,

Sergio

Cholera toxin subunit B conjugated with Alexa Fluor 488 or 594 (Invitrogen-Molecular Probes, Eugene, OR, USA) is a proper choice. 3 days after injection is OK for observing the motor neurons in the spinal ventral horn.

Hi Anya Snary ,

I agree with Martin Schol , I use Superfrost Plus™ Slides by Thermo Scientific which electro-statically attract frozen tissue sections. I mount with them directly from sectioning in the cryostat and the tissue

is adhered to it properly.

Good Luck!