- Susan Bachus added an answer:1Does anybody here have a softcopy of A Handbook of Normal Penaeid Shrimp Histology?
I'm working on penaeid histology and I don't have accesses to Appropriate and standard reference.does anybody have a colorful copy of Bell and Lightner book(pdf format)? I would appreciate to suggest any other standard reference.Following
- John T Garretson added an answer:1Specific molecular marker for ANS fibers?
I was wondering if anyone know the specific or unique molecular markers for the following ANS fibers (I would like to do some immunohistology to distinguish between the non-ANS nerve fibers and the ANS nerve fibers, and between the ANS nerve fibers):
1) sympathetic fibers
2) parasympathetic fibers
3) enteric fibers
Thank you very much in advance.
In what tissue are you trying to distinguish ANS fibers from the background? Brain? Peripheral NS? In which organ? It might help others assist you if you are more specific.
- Christophe Cisarovsky added an answer:6Does anyone have a clue for a nice TMA DAB staining?
I have the following problem:
I'm doing a usual DAB staining (1°Ab, 2° Ab conjugated, HRP and Tyramide and then DAB for chromogen detection) that works well.
My issue is that many of the TMA samples fell off the slide during staining procedure as well as I have many "DAB dirt" accumulating on top of my slides (I use slide racks with plastic slide holders) and thus preventing optimal penetration of the DAB and efficient washing afterwards.
I do deparaffinzation 10min at 60°C followed by 3x Xylen, 3x 100% EtOH, 1x 90% EtOH, 1x 70%EtOH, 2x dH2O (each 1min).
Dos anyone have any advices for me?
Thanks in advance!
Thank you all for the suggestion. I will try them!
- Akhilesh Kumar added an answer:3Any advice on staining after using Bouin fixative?
We fixed our specimens(rat testis) by Bouin's fixative.when we stained the sections by H&E and Toluidine blue but the slide not found suitable staining what can I do to resolve this problem?
thanks , Muss sir, for fixative. i was also use DAVIDSON'S and formalin, both of these was better for testis staining (H/E).Following
- Nalini Poojary added an answer:12Any suggestions on removing DPX mountant from fixed slides?
I mounted DAB stained sections onto a slide and used DPX as a mountant. There seems to be some sort of a problem in some of these sections post mounting. I would like to remove DPX and re-mount the sections. Has anyone done this previously? Is there a standardized protocol for this? A few important points :-
1. The sections were not dehydrated with ethanol dilutions prior to mounting.
2. The slides are not old. They were mounted less than a month back.
3. The immuno has worked in most of the sections/slides. Its only some slides that are problematic.
Yes, I agree with Gudrun Lang. But take care to see that the section is not washed off the slide.Following
- Ceara Mcgowan added an answer:12Any advice on methods for double embedding small tissue (agar and paraffin)?
I am trying to double-embed very small tissue pieces for histology and am having some trouble. The tissue is tiny and in order to orient it properly (and not lose it) I have been embedding it in agar (1 or 2% low melting point) and then dehydrating as usual. My problem is that the paraffin is not infiltrating the agar and I get a soft blob in the paraffin block that is not stable enough to cut. I have tried expanding the length of time in dehydration and clearing, but that hasn't worked so far. I also refreshed my ethanol so I am sure of the concentrations. I only expanded the time in each ethanol solution by about ten minutes so far. My procedure looked as follows:
Tissues were fixed in formalin. After three (sometimes more) days, tissues were embedded in agar and then placed in formalin overnight.
Tissues then underwent dehydration:
- 70% EtOH 30 minutes
- 70% EtOH 30 minutes
- 80% EtOH 30 minutes
- 95% EtOH 30 minutes
- 95% EtOH 45 minutes
- 100% EtOH 45 minutes
- histochoice clearing agent 30 minutes
- histochoice clearing agent 40 minutes
- histochoice clearing agent 40 minutes
- paraffin 30 minutes
- paraffin 40 minutes
- paraffin 45 minutes
- embed in paraffin blocks immediately
The agar blocks are still soft after this procedure. The protocols I have read mostly state to "proceed with dehydration as usual" after embedding in agar, but that is not working for me at all. Help would be greatly appreciated!
We use the double embedding of agar and paraffin process and it work great. We fix our tissue and embed in 4% agar, process overnight and then embed in paraffin, I think if you increase your agar from 2% to 4% you wont have a problemFollowing
- Mohamed A. A. Mahdy added an answer:7Wich protocol are you using to stain frozen liver sections with oil red o ?
I have tried till today a lot of protocols (IHC world,...), using isopropanol or propylene glycol or both, but I haven't had any satisfaying results yet...The stain is scattering everywhere out of lipid droplets, red oil is precipitating and the slides end up all dirty. The last protocol I have tried is the following one: fix in formol 10% for ten minutes, then put in 100% propylene glycol for 5 minutes, stain with oil red O 1,8% in isopropanol ( dilution from stock solution 6:4) and then put in 85% propylene glycol for 2 minutes and rinse in deionized water for 1 minutes. I have also tried to later stain with Gill III hematoxylin for 30 seconds, but the results were even worse... I am quite desperate about it ! Please if anyone met these kind of problems and managed to solve them, I would really appreciate her or his help :) Thanks in advance !
I stained skeletal muscle cryosection (10 um) with ORO using Isopropanol and works fine
You can try this protocol
- Anudeep Venkata added an answer:4Why is acetic acid crystals seen in Benzidene staining?
Help me to get rid of acetic acid crystals in Benzidene staining??
During benzidene staining of haematopoietic colonies there are lot of crystals i can see and after washing once with PBS only few crystals are seen and how to get rid of those acetic acid crystals?
Dear Gurdun Laang,
Thanks for valuable suggestion.Following
- Wilfredo Molina Wills added an answer:6Any advice for a nucleolus staining?
I have samples that were prepared for TEM (fixed with formaldehyde and glutarldhyde, post fixed with osmium and embedded in spur resin). I would like to take some semi-thins section (0.5 micron), stain them with a nucleolus staining and then imaged them in light microscope.
Any suggestion, to which staining I can use ?
I think that immunofluorescence techniques and fluorescence microscopy, including laser confocal microscopy, it is the best tool to detect and analize the intracellular location of various cell components. Fluorescein or rhodamine have been commonly used for immunofluorescence microscopy. A variety of nucleic acid binding dyes have been developed. We can use this method for histochemical staining and observation by laser confocal fluorescence microscopy.Following
- Senka Pantic added an answer:18How to remove coverslips from mounted slides ?
I immunostained whole mesenteries from mouse. I mounted them in Dako Fluorescent Mounting Medium. Now I would need to counterstain them. Does anybody has any idea to remove the coverslips from the slides, in keeping the samples safe ?
Thanks a lot !
Put slide into the cuvette with xylol for 60 minutes.Following
- Michael Scholz added an answer:6Any suggestions on 3D tissue reconstruction software?
Hello, we are working on different ways to digitally reconstruct histological tissue sections with a very high accuracy for anatomical purpose. Has anyone good results with a specific and commercially available reconstruction software?
Thanks a lot for your valuable and helpful hints and comments.Following
- Ernest R Schockaert added an answer:14How long should I fix in Bouins for Masson's Trichrome?
Our current protocol has us fixing slides (cryo or paraffin) for 24 hours. It was just brought to my attention that this is wrong, but I was not told how long is correct. I would love to know what others do for their protocol.
How long are you fixing the slides in Bouins Mordant before doing a masson's trichrome stain?
As some colleagues already said, the time of fixation depends on the the sample (and mainly on its thickness). Bouin has the great advantage that fixation time is not very critical (even though it is not the best fixative on the cytological level). I always have used Bouin in field conditions, when collecting free-living flatworms of around 2mm in size (or less). I leave them in the fixative (in an embryo dish) until I have the time to transfer them to alcohol for later processing (embedding and sectioning) in the lab. When I am just a few days away, I even leave them in the Bouin! In the lab I prefer a sublimate- and formaline based fixative, when I have controle on fixation time. Bouin also has the advantage that the sections take the dyes very well (such as Masson's of iron hematoxylin). But the time one leaves the sections in "the mordant" depends on what you want to see and asks for some experimenting. So all various times which have been mentioned so far (including yours) are correct when you are satisfied with the result ;-). One last thing: do not use Bouin that is too old! You better add the acetic acid shortly before fixing. Good luck, Tanya!Following
- María Clara Carou added an answer:18Does anyone know a histological image analysis program?
I'm looking for an image analysis program in which I can make some measurement in histologic sections of cartilage. I am especially interested in a program which will make it possible to give surface area information on a manually selected area. Also drawing lines, to divide it into three equal parts would be nice. Does anyone know such a program? Photoshop doesn't have these tools unfortunately.
Leyca Q Win; Image J.Following
- James Clinton Shawulu added an answer:11Why does my spinal cord sections shred when mounted and left to dry?
I´m working with 20 microns sections and whenever I mount them and leave them to dry, they shred. If I don´t let them dry, they´re fine, but they don´t stick to the slide, falling off in the staining process. I´m losing a lot of samples and I´m trying my best to find the perfect timing between mounting and staining, but nothing seems to work. Does anyone already worked with this and had the same problem?
I do not know which fixative you used. However, if they are well fixed they will not fall off. Secondly, you should after sectioning free air dry your slides and prefix them in hot oven at 60°C before staining process. Use superfrost slides they are good. Depending on the period you want to stain, they should be in the oven. The thickness matters about what you are studying the tissues for, maybe white matter gray matter architectures. But if you are looking for cellular architectures, you should have thin sections. Again the use of H & E is good. Currently i am doing some and they are beautiful.Following
- James Clinton Shawulu added an answer:6Method for harvesting newborn mouse and embryo brain and spinal cord?
I want to do immunohistochemical analysis on newborn mouse and 20 days embryos. I harvested brain and spinal cords of 90 days mice but its so different with newborn and embryo. Is there anybody who have experience with newborn and embryo? How I can do immunohistochemical analysis on newborn and embryo? Is it necessary to harvest spinal cord from back bones or we can directly use spinal cord with bones attached them for 20 days embryo? Is there any protocol?
Since your targets are the nervous tissues, i suggest you fix your tissues with fixative for nervous tissues when you have removed as much flesh or bones as you can. That will make the brain or the spinal cord fix well. Do not care much about the left over muscles or bones since they are not your target. With correct fixatives, it may not affect your result.Following
- Arash Khodamoradi added an answer:9Is there an alternative way to adhere 150 micron golgi-stained hippocampal sections to slides?
We have been using the superGolgi Kit by Bioenno Tech and have had two major issues:
1. Preventing sections from fragmenting due to drying out.
2. Having them adhere to our slides.
The first problem is directly related to the second problem in that any time we have tried to stain them after the mounting step, they have fallen off of the slides when placed in 0.01 M PBS-T. Therefore, we need them to dry a bit more for them to stick properly, yet in doing that they have a greater chance of drying too much and fragmenting. So, we were wondering if using a different, possibly more adhesive, slide might help. We are currently using 1% gelatin coated superfrost slides and were considering increasing that to 1.5% (2% did not seem to work as the gelatin would not disentigrate). Basically, if anyone has any tips on how to help them adhere to the slides and not dry out, it would be greatly appreciated!
Hi Dear Researcher
Unfortunately I have no experience in this field.Following
- Wolfgang H. Muss added an answer:6How do I perform a chrome alum hematoxylene phloxine staining?
i was following an old protocol for chrome alum staining. standardizing the procedure was finding difficult.1.tissue comes out from the slide while phloxine treatment 2. no uniform staining 3. chrome alum stain is some times not working ##please help me to prepare good solutions for the staining and give me some more references .
Dear Rajesh, I only can second the opinion and fact finding of Gudrun. I personally emphasize - if you are reproducing a method or technique first time - to follow a given original recipe step by step, using all the solutions and chemicals given there. "Modifications" in/of solutions and chemical always may have some impact on outcome, yielding sometimes wrong / false results. Losing / loosing tissue from the section may have to do with the strong acidic pH of the Harris' Hematoxyline, as Gudrun pointed out, but may have also other causes: Unfortunately you didn't tell about the tissue you are staining (and more interestingly, how you processed that tissue before staining...i. e. fixation, processing into paraffin etc.). Good luck and best wishes, Wolfgang.Following
- Milena Pesic added an answer:8Why during sectioning of undecalcified plastic embedded bone, sections start to tear in region of secondary spongiosa?
I would highly appreciate if someone could give me an advice regarding plastic embedding. I am working with the bone tissues, and I am trying to establish embedding process in Spurr resin in order to analyse dynamic histomorphometry. Bones are from 6 months old mice. First, I dehydrate them in 70%, 95%, 100% ethanol and 100% acetone (every step takes 2 days) and then start infiltration process with Spurr resin 50%, 75% (mixed with acetone) and 100% (also steps take 2 days). Dehydration and infiltration processes take place in desiccator connected to vacuum pump. At the end of infiltration, bones are embedded in 100% Spurr and plastic blocks polymerize at 55oC for 2 days. When I do sectioning, I have this problem that region of secondary spongiosa starts to tear, and also block by itself is very brittle. What could cause this problem, could it be that infiltration was poor, or the ratio of Spurr components are not good?
Thank you for any answer and help,
I am working now with MMA, and it works great. Sections look very nice. Thank you all for your answers and help.Following
- Mohammad Nasir Uddin added an answer:5How can I prevent detachment of retina from choroid during cryostat sectioning of human optic nerve head?
Hi, in my project I am doing cryostat sectioning (Longitudinal section) of human optic nerve head. In cryosection, I have to keep all the three layers (retina, choroid, sclera) attached together but after cryosectioning I find the retina detached from choroid or huge gap between retina and choroid. I think this problem might be due to embedding problem because when I embed the tissue in OCT medium I can’t keep the retina completely attached to choroid. It would be highly appreciated if you have any suggestion to solve this problem.
Thanks everyone for your suggestions. Definitely it helps. However, I want to share my protocol briefly so that you can have better idea about my problem and suggest accordingly.
Yes, it would be better if we could fix the eye before the cornea and lens removal as it allow the retina to remain flat and attached to choroid but the problem is most of the time we receive the eye from eye bank without cornea and lens as cornea is kept for transplantation. So after receiving the eye or better to say eyecup, I fix it in 4% PFA for 2 hours before application of 10%, 20%, 30% sucrose solution for cryopreservation.
Another fact that I must share is, I don't embed the whole eye cup in OCT medium. My particular interest is ONH and the area surrounding the optic nerve head. So, I cut a rectangle shaped area (e.g. 12 mm X 8 mm) keeping the ONH in middle. Then I embed the tissue horizontally in OCT media facing the retina up. As soon as I place the tissue in the OCT medium the retina tend to lift off the choroid a little bit which I can't prevent. Then I do the snap freezing in Isopentane kept in dry ice.
In cryosectioning step, I use the object temperature -20 and section thickness 10 micron. After cryosectioning when I check the sections under microscope I see a huge gap between the layers (retina, RPE, choroid) and I think the gap is justified because it already had gap between retina and choroid during embedding process.
This is the details of my protocol and problem. I badly need to solve this problem. Therefore, I am looking forward to your suggestions. Thanks again for your concern and valuable feedback.Following
- Mohamed Najimi added an answer:13Storage of tissue for immunohistochemistry?I am going to perform a mice experiment abroad and plan to harvest tissue (adipose tissue, liver) for subsequent histology and IHC. After one week I will return home. I know that fixing of tissue in 10% Formalin of more than 24 hrs may affect further IHC. I am grateful for advice about the best storage/transport medium after fixing?
It's possible but you have to made many improvements to your tissue before IHC for antigen retrieval.Following
- Khalid Eldahan added an answer:8How can I label brain hemispheres for free-floating histology?
I am running immunohistochemistry on rat brain tissue, and the sections will be stained while free-floating. However, we need to reliably know the left from right hemisphere. We cannot take a notch out of the cortex (which we have done previously), because we are taking a full survey of the brain.
Could someone recommend a method to label the hemispheres? Is there some sort of marker or pen that is visible to the eye but would not affect staining?
- Thank you -
I would use a little dab of lipophilic dye, like DiI:
Are you planning to use fluorescent IHC? If you do, just make sure that the dye doesn't overlap with the emission profile of the fluorophore you will use!Following
- Chang Yi-Fen added an answer:2Does anyone have a protocol for revealing BDA tracing in the brain with a Streptavidin Alexa Fluor conjugate?I am wanting to use BDA for neuronal tracing and rather than revealing this with DAB I would like to use a fluorescent tag, for viewing on the confocal microscope.
I also tried to tracing neuronal circuit by BDA conjugared HRP. I have a question about "blocking". Due to HRP, do I need to use 3% H2O2 to decrease endogenous peroxidase?
Please Help me, I have tried for this over 3 month. Thank you very muck.Following
- Virginia Gutiérrez added an answer:9What is the pattern of Nitrotyrosine staining in normal and injury induced mouse liver?
I am using oxidative stress marker Nitrotyrosine IHC staining on mouse liver. I am getting desirable results but I have also observed positive sinusoidal staining in normal control mouse liver. Has anyone noticed a similar issue?
When you have mouse on mouse ihc it´s normal to have unspecific staining in the sinusoid, it means that the blockade has not worked properly.I recommend you a blockade with a anti-mouse FAB fragment (you can obtain it from Jackson immunoresearch) for 1h at RT at 1:10 concentration.
Hope this hepls
- Vera Weisbecker added an answer:4Are there any tips for MAB377 which is not working with formalin-fixed mouse brain nuclei?
The tissue is fixed in 4% PFA/PBS (no methanol) and ground up for approx. 5 min. in 1% Triton-X 100 and 40 mM Sodium Citrate for subsequent flow cytometry counting of free nuclei - then washed and stained as per usual (2 washes after the primary stain. I’ve used different incubation times, temperatures, PBS types, secondaries (Alexa Fluor 647 and 488), freshly fixed (1-2days) and older brains, nothing works. I use up to 1:10 dilutions of MAB377 in a solution of roughly 10^6 cells/ml. If anything, the nuclei are darker than the tissue debris under the fluorescence microscope, it is as if nothing gets in. I have used two batches of MAB377 (Merck Millipore) – they seem to have both been frozen (the latter batch very briefly on its way to me) but surely that won’t totally destroy the antibody? Any ideas or alternatives?
Hi All, thanks for your responses. As an update, it may have been the fixative indeed - the protocol seems to work with specimens fixed in formalin that was _first_ dissolved in distilled water, and had 10x saline added afterwards (I had previously cooked up the PFA in saline, which seems to diminish its fixation abilities according to a histologist I talked to). But I still need to confirm that this really is the main problem. The fixation times are really long - weeks up to 1 year - and fixation in the "wrong" formalin seems to very quickly degrade the ability of the sample to stain. Hopefully I'll figure it out soon and I'll post a final solution then :-).Following
- Hamid Marvasti added an answer:11What is the protocol for Wright staining ?
Can anybody help me to get the protocol for wright staining? I want to stain the different cell present in sputum.
Dose any body have wright Giemsa SOP for cell (tissue) staining please ,Sigma productFollowing
- Maja Løvbakke added an answer:4How do I treat tissue for paraffin embedding after its been fixated in Carnoy's Solution?
I wan to look at the mucus thickness and Carnoy's solution have been used as fixative for this purpose before.
From what I can read, it is seems common to fixate between 2-4 hours in Carnoy's solution, then transfer the tissue directly to 100% alcohol for 24 hours. Some seem to use shorter time in alcohol and use methyl benzoate for 24 hours before xylene dehydration. I am confused about what give the best result or give the least shrinking of the tissue.
Thank you very much for your help everybody!Following
- Sandra Franz-Guess added an answer:4How do I stain using fuchsine basic?
My colleague would like to stain Branchiostoma using Fuchsine basic. Does anyone have experience with this technique? We're looking for details on concentrations since we can't really find anything useful on the internet. Everyone seems to use it as an additional staining agent, but not as sole agent. I'd be grateful for any help you can give us!
Thank you for all your input! My colleague looks for an overall stain, so no specific tissues that need to be highlighted. I'll forward your tips, I'm sure it'll help them. Thanks!Following
- Yogesh Kanna Sathyamoorthy added an answer:4Can anyone provide me with a method for conducting histological investigations, I wish to study Ischemic events in neonatal rats?
But I could only find developing mouse brain atlas. Can anyone recommend a single developing rat brain atlas that is available for download?
thank you Dr. HousmanFollowing
- Closed account added an answer:11How do you store tissues before frozen sectioning?
Fairly soon I will be finished decalcifying my tissues for frozen section preparation. Normally I would then pass the tissue along to my lab specialist to perform frozen sectioning. However, that specialist will be leaving soon, and I want to make sure my method for storage works until another cutting specialist arrives.
I plan to wash the tissue with 1X PBS multiple times and cryopreserve the tissue in a 30% sucrose solution at 4 C overnight. The next day I will embed the tissues in OCT compound and store at -40 C in the deep freezer. Would this method work in preserving the tissues and their possible fluorescence? Thank you very much!
I store both tissue and cut sections in -80. If kept in the dark, it should retain its fluorescence.Following
The study of the microscopic anatomy of cells and tissues of plants and animals, including tissue fixing, fixation and staining.