- Monika von Düring (Duering) added an answer:5Fixation of acute brain slice...
I want to fix acute brain slice after cutting and want to do immune and histology. My question how will I fix slices for these experiments? Can anyone here for giving advice?
Thanks in advance
we use 4 % PFA in PBS for 2 hours at room temperature on a shaking device - best wishes MonikaFollowing
- Leslie Rietveld added an answer:7Anyone have an automated method for counting cells/determining cell density on stained brain slices?I have cresyl violet stained brain sections and would like to look at cell density in different regions of the brain including the hippocampus and areas of the motor cortex. In some places the cells are too densely packed to be easily counted. Has anyone used image J and regions of interest to quantify cell density in brain?
When I used cell profiler I did not use any pre-made pipelines. I created my own based on my particular needs. However I was able to find a link to these pipeline examples that you can download and test out.
- Hannah Bader added an answer:3How can I avoid from autofluorescence in mouse liver tissues?
I am using mouse liver tissue which is fixed in 4% PFA and embedded in OCT. However, autofluorescence signal is coming from the negative control samples, when Abcam protocol is used. Do you have any idea or any good protocol that could work with these conditions?
autofluorescence is usually only strong in the green channel (FITC, Alexa488), but not in the red (Rhodamine, Alexa 5XX etc.). If you do not do a co-stain, switching to red may solve the problem. Also, often you can distinguish signal from noise by comparing your stained section with secondary only control. If you have a decent signal, it will be considerably stronger than the autofluorescence. In that case, you just have to adjust the exposure time so the camera can see the signal, but not the noise.
- Sophia Ekeuku added an answer:3Could anyone recommend an efficient protocol for histology of osteoblast cell, adipose and liver tissue culture and also muscle tissue?
Could anyone recommend an efficient protocol for histology of osteoblast cell, adipose and liver tissue culture and also muscle tissue?gy
I want to do H&E staining on a cell tissue culture. If i can't embed and section how do i go about it.Following
- Maria Mendoza added an answer:3Anyone can help me please..Carbohydrate dye?
I don't have any idea about dyes >> so If anyone can help me..
I need dye for only carbohydrate in microalgae ..
PAS and you can review the book
Pearse AGE. 1985. Histochemistry: Theoretical and Applied. Vol. 2: Analytical Technology. Edinburgh: Churchill-Livingstone. 1055 p.Following
- Wincenty Kilarski added an answer:10What is the best method to stain bile acids in histology sections?
I wonder how to stain for bile acids in tissues?
sorry no more sugestions
- Ramzi Al-horani added an answer:11What are the best intra-cellular markers for regional intracellular hypoxia/ischemia?I am trying to describe hypoxia in the cell core comparing to the cell surface. Traditional markers like HIF-1 are mostly measured in whole tissue not single cells
You may think about the level of O2 reduction to H2O at the level of electron transport chain in mitochondria. I think this can be measured through measuring the activity of the enzymes of that chain. at the surface of the cell, meaning cyctosol, give a glimpse if oxygenated myoglobin can assist for your purposeFollowing
- Sebastian Schmitt added an answer:2Can anybody guide me the protocol to get better tissues of sciatic nerve for histology?
I am working on EAN Model, to perform histology i need very good slices, but unfortunately i couldn't get proper slices, tissues get distorted when we stain, we have made all possible changes but couldn't proper tissue for histology. any body doing HE or Immunostaining of sciatic nereve could guide me, Thanks.Following
- Héctor Osorio-Vega asked a question:OpenCan i detect a single cell mutation in a tissue?
Is there any technique associated to histology that allows to detect a mutation in a specific gene of a single cell of a certain tissue? (i dont need to know the type or detail of the mutation, only that there is one), assuming that the rest of the cells have a wild type geneFollowing
- k.a Galil added an answer:7Can anyone help with cryosectioning of mouse liver tissue?
I just would like to know a few general tips - What thickness should sections be? What temperature should the cryostat be? Does orientation of the tissue on the chuck matter? Would you recommend a particular lobe or area of the liver to use? Should I leave the sections to air dry once they're on the slide (How long? At room temperature or 37°C?)? Can the slides be stored for long once the sections are on them (how long? at what temperature?)?
I intend to perform H and E, as well as Oil Red O staining on the sections.
That's a lot of questions, I know! But any advice at all would be greatly appreciated.
hello: i agree with many of my colleagues but I think paraffin sections will let you control the situation better and you get good results when you stain either H&E or even Histochemical stains. you obtain a good morphology and preserve the slides for further examination.
one thing about research is many times i have gone back to my old slides and saw items i missed the first time.'
i know croysections are faster and quick but if you intend on through examination i feel paraffin sections are better. As for orientation yes it is very important to orient to the direction of the lobe you want to examine ,i usually draw on a piece of paper the orientation of the liver and the lobe and keep it in the file believe me if you don't have orientation many lobes look alike and one tends to blame himself by saying i wish i know which lobe is this??
I use professor john kiernan’s book on histochemistry he is also my colleague in my department and a well respected histologist and histochemist in the world
You can see all his achievements here http://publish.uwo.ca/~jkiernan/aboutjk.htm and his new book
Kiernan, J. A. 2015. Histological and Histochemical Methods: Theory and Practice. 5th edition. Banbury, UK: Scion Publishing. (ISBN 9781907904325). has just been published and is available
good luck and best wishes
Professor K.Galil.DDS.,D.Oral&max fac Surg.,Ph.D.,FAGD.,FADI.,Cert.Periodontist (Royal College of Dental Surgeons)
Patenet developer of < DGOHE +itune>website , visit to see the new Oral histology App for the I phone,Ipad &I Pod touchhttps://itunes.apple.com/us/app/dgohe+/id1015383607?mt=8Following
- Prashant Battepati added an answer:5Can stereomicroscopy be used to study dentinal tubule morphology?
As scanning microscopy is comparatively expensive, can stereomicroscopy be used to analyse dentinal tubule morphology ( I agree and understand the detailing is best under SEM) if yes what magnification is ideal..
Thanks a Ton Max for this wonderful explanation!...,Following
- Daniel Lerda added an answer:2A FISH protocol (DNA beacons for Epifluor) says denaturation @ “150 deg C”. Is this too hot?
I have 2 protocols for FISH w/ DNA beacons (approximately 25 nucleotides long) for use on formalin-fixed paraffin embedded tissue (5µm samples) for examination with epifluorescence.
One protocol says to heat the tissue slides to “150 deg C” and then to cool to slides to 90 deg C, layer beacon probe over tissue of interest. Then lock the temperature at the melting temp of the beacon (usually 50-70 deg, but no specific stem melting temps given) and hybridization time 20 min. Bring back to RT. Do 3 consecutive washes in distilled H20. Air dry and examine under Epifluor.
The other protocol says to layer beacon probe onto tissue area of interest and to cover with hybridization slips and serially heat (no increments given) to 90 deg C for 10 min and then lower temp to 70 deg C for 15 minutes. Bring back to RT. Do 3 consecutive washes in PBS pH 7.0. Air dry and examine under Epifluor.
Questions: The second protocol's denaturation temp looks more reasonable. Would second protocol be better? What would be the increments of serial heating? Can the washes be with distilled H2O instead of PBS?
Dear Jennifer Souders,
The second protocol is the best and with respect to PBS, I consider it appropriate. Best regardsFollowing
- Stefan-Claudiu Mirescu added an answer:3Dear collegues, do you have access to any articles or knowledge concerning the golden ratio (phi) in the human anatomy or histology?
Many articles have been written concerning the presence of the golden number or the numbers of Fibonacci in architecture or botanical sciences. Are there any publications in the field of human anatomy or histology? Thank you!
Thank you, dear collegues, for your responses!Following
- Diego Yoshikay added an answer:2How can I keep morphology using OCT or Crymatrix in Plant vascular cambium cells?
Im working with fresh apical stems and this organ is very soft. After embed in Cryomatrix or OCT, it is posible to obtain around 40% of sections with good morphology (even cambium cells). But in the other 60%, shrinkage happens specially in cambium cells. Troubles:
- Samples after dry suffer of cell shrinkage, specially the cambium cells
- Samples after ethanol wash (for laser microdissection), suffer an even more shrinkage.
How could I avoid shrinkage in my cells?
Nop, ethanol is used to dry sections and fix before laser microdissectiion
Could be changed by acetoneFollowing
- Glenn Sherer added an answer:5Does interstitial growth of cartilage give rise to isogenic groups of chondrocytes as a final aim?
Or it´s only transitional stage of chondrocytes formation which is separated into a individual cell of chondrocyte later? And why should it separate, whereas it differentiates from a single chondroblast? Couldn´t it just differentiate from a single chondroblast?
I mean, if it´s give rise to isogenic group, what´s the purpose of this formation? What it benefits? Why not stay still in form of an individual cell?
Sorry to keep asking such basic stuffs that an undergraduate should´ve already known well.
Several others, above, have responded to your question from the perspective of "how" cartilage differentiates and grows -- but not "why". I'd like, instead, to comment on the assumptions that seem to underlie the way you chose to articulate your question, i.e., that biological structure and function are consequences of design focused on serving a predetermined purpose.
Needs and purposes cannot determine their own means of fulfillment. Such teleological thinking was abandoned by biologists very long ago. The only meaningful answer to "why ... ?" in biology is evolution. Cartilage grows the way it does, and has the form it does, because it evolved that way and because the consequence of that evolution served a useful (beneficial) purpose. Evolutionary change is a consequence of selection exercised on the consequences of random events (mutations). Cartilage and all other tissues are the way they are not because those ways best serve a purpose of design but because once they emerged they persisted, and all that is needed for evolutionary changes to persist is that they contribute to the success of the species of organisms in which they emerged. Emerged (evolved) solutions need not be (or have been) the best possible ones.Following
- Henry Lopez added an answer:2Does anyone know how much hydroyproline is in a normal size mouse lung- what about one challenged with bleomycin?
Several publication have different values of hydroyxproline in mouse lungs with and with out being challenged with bleomycin. In my a recent study I ran, the values were similar between naïve and bleo challenged mice. In a cohort study the Mason trichrome histology showed huge differences. Ran the study according to vendor.
Thank you in Advance,
Thanks Willi- I feel much better quantifying the lungs histologically.Following
- Paul Jason Thomas added an answer:9Any suggestions about the presence of breakage and holes in cardiac frozen tissue sectioning?
I hope all of you have a great good day.
Lately I have severe problem with my mouse heart frozen tissue sectioning.
Here my protocol :
1. after collection of the heart organ, I wash with PBS and preserve with 4% parafolmaldehyde (in PBS) for 2-3 days in 4 degree Celcius.
2. Soak in 20% sucrose for 3 days until the organ not floating anymore (some of sample is almost 1 weeks).
3. Dry with tissue towel and put in OCT compound for 30 minutes, following with methylbutane freezing method .
4. Tissue sectioning, 8 micron thick and do the H&E staining.
And the problem is :
1. After sectioning, before H&E staining, I check the slide and the breakages (holes) are presents. The morphology was so bad.
2. Then I realize, directly after the sectioning, the breakage was not occur, but around 1 minutes after I put the slide under the microscope, the breakage getting bigger and bigger and ruin the morphology.
What I already tried :
1. Make the section thinner and thicker did not really help.
2. Freezing technique using methylibutane-liquid nitrogen also did not help ( quick freezing to avoid ice crystal artifact)
3. After sectioning, directly put the slide in to PBS contain PFA also nor work.
So I really don't know what really the problem is.
I would really appreciate for your suggestion.
Thank you so much for the help.
We do not fix the heart at all. Just place in PBS for 30min, gently tease out any blood clots and dry excess PBS, then place in OCT in an embedding mold for 30-45min. Then, freeze for 20-30min in dry ice cooled isopentane.
Have you tried a different set of slides? My guess is that the tissue is not adhering to the slides properly, and therefore the morphology is distorted when the sections dry on and pull away from the slide. You could try coating your own slides (a pain) or just a different batch of commercial slides. We use Fisher Colorfrost Plus slides. It also helps to warm the slides before adhering the sections to them.
- Guillaume Gauchotte added an answer:3How can I analyze collagen fibers on mouse skin that is stained with trichrome?
I want to compare two below images for example.i want to know, in which image exist more collagen fiber? for this goal, i need to know, how can use Image j software, step by step, because i never use this software.
thank you for your attention.
If possible for you to perform another staining, I think that a staining with Sirius red would be more reliable to quantify collagen by image analysis.
If you have Adobe Photoshop, it is very quick and easy to evaluate the collagen area with the "color range selection tool" (then divide the number of selectid pixel by the total number of pixels of your picture).
- Sumana Chakravarty added an answer:6Any 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?
As per my experience, Bouin's is ok for testis fixation. I got very good results in PAS+ stain and H&E as well. Only thing is you have to cut it in pieces like transverse or longitudinal (tubular arrangements will look same in both) for better penetration say about 24 hours of incubation. Freshly prepared fixative is always better. Regarding yellow colour (picric acid) of the tissue, don't worry about that. During the staining process of dehydration using graded alcohol series eventually it will go away.
All the bestFollowing
- Timothy J Seabrook added an answer:5In freezing brain tissue for WB - do you need sucrose?
In my current lab they dump freshly isolated brain tissue into 0.4 M sucrose before freezing on dry ice. Is the sucrose necessary at all when I only want to do WBs i.e. could you get equivalent results by leaving it out? I want to detect protein phosphorylation so I strive to keep the tissue frozen as long as possible before synaptosome preparation and with sucrose I need to thaw the tissue more then if I'd leave it out.
I have run WBs, including for protein phosphorylation, using brain tissue directly frozen in liquid nitrogen. If a crude protein isolate, with subsequent WB, is all you plan to do than sucrose is not necessary. However, if you want to look at proteins in synaptosomes, then sucrose is likely required.Following
- John K Fellman added an answer:4Is there a replacement for Meldola's Blue Dye with respect to PCN egg staining?
I am having trouble sourcing Meldola's Blue Dye in the UK and the only supplier I can find has discontinued their product.
Is there an alternative dye which can be used for staining when determining viable/nonviable PCN eggs?
Just found this, too! 5 grams for $255.30USD
- Hadi Taghizadeh added an answer:4How can I fix stretched tissue and be sure to fully capture the tissue in that stretched state?I'm going to inspect microstructural changes induced in the arterial wall as it is stretched in the physiological pressures. I'm using tissues extracted from aorta and preserved in PBS.
I need to fully fix tissue samples in that stretched state. Are there any protocols for this?
Main parameters to include in our investigation is the dimensional changes in the layers and cells of micrometer size. Then desired fixation method should fully maintain stretched state.
Any contribution is highly appreciated.
Thanks for the interesting responses, I've tried something similar to the method proposed by Robert, however the results were not satisfactory. As David stated, the crosslinks are not well preserved with formalin.
By the way, I'm extremely interested in quantification of the collagen fibers within soft bio tissues.
I've started a new research on the tendons and ligaments and trying to identify the interaction of the collagen fibers with mineral molecules in the interface of the these soft tissues with the bone. any suggestions?Following
- Yasushi Nakagawa added an answer:4Any solution for uneven EdU labeling on mouse brain sections?
We are using the regular Click-It kit to detect EdU on mouse brain sections, and are having a problem of uneven staining. In many cases, only one side of the brain or only some part of the brain has labeled cells. Does anyone have a solution for this issue? Would it help to dissolve AF azide in PBS instead of DMSO?
Thank you for all the suggestions. We are doing a short-term pulse labeling with E14.5 to E16.5 mouse cortex (with 2h-18h survival time). We have 15-20 small sections on each slide and permealize the sections for 20 min in 0.5% Triton X-100/PBS. We have tried to use coverslips (with 150ul of reaction solution per slide) or without coverslips (250ul per slide). We are using 0.6ul or 0.3ul of Alexa 594 or Alexa 488 azide per slide (Invitrongen, dissolved in 70ul DMSO). It seems it's worth trying to increase the volume by a lot. If it does not help, we will try a longer permealization. Another thread started by Justin Judd suggests other vendors (Lumiprobe and Carbosynth) have much better prices for dye azide (they are actually water-soluble) and EdU, so we will try those, too.Following
- Kevin C Chen added an answer:3Which is the best blade inclination to cut very healthy coronal hippocampal slices?
i have some questions for you. I'm trying to performe fEPSP experiments to induce LTP in hippocampal CA1, but i have some problems finding a good response. I would to record fEPSPs from the dendritic zone of the CA1. I'm thinking that one of the problems could be the slicing angle. Which is the best angle to cut healthy coronal hippocampal slices? Another question is about the stimulating electrode. I usually use a glass pipette filled with acsf to stimulate the fibers . Is a good choice or i could use best stimulating electrodes?
Thanks so much for help ;)
Hi, when you said you want to cut "coronal" hippocampal slices, I assumed you meant the dorsal (the upper part) hippocampus. Usually, a normal coronal cut will do (that means cutting is perpendicular to the mid-line). Since you are not getting good fEPSP responses, you may want to tilt 10 degrees along the perpendicular line.
As to the stimulating electrode, a glass pipette (similar to a mono-polar electrode) is not a good idea. I think it is not the resistance issue, but that a mono-polar stimulating electrode is affecting too many fibers bundles (work out the source-sink circuit for mono-polar electrode yourself). Most people use bipolar stimulating electrode, which will only stimulate smaller range of fibers. A better one is a concentric type, commercially available. But you can also make one yourself, by twisting two insulated wires together, and remove the insulator at the tipsFollowing
- Michael Gudo added an answer:12Why my human cartilage histology wax sections are broken and empty?
I have sectioned some wax blocks today using the Microtome but my sections are broken and some are empty. Any ideas why that is? I followed all the steps in the protocol as it was stated.
samples: Human femoral head cartilage round plugs, dimensions 10mm diameter, 2mm depth
Fixation - 10% NBF for 48 hours
Decalcification- 0.5N HCL/0.1%glutaraldehyde for 48 hours
(left in 70% ethanol 4C fridge , over the weekend)
Dehydration- 70% ethanol 1 hour, 90% 1 hour, 95% 1 hour, 3x 100% 1 hour
Clearing-histoclear over night, histoclear 3 hours (left in 70% ethanol overnight)
Wax- paraffin 3x2 hours at 60C
Any help would be appreciated.
The glutaraldehyd step is also not necessary in my opinion, since this fixation is for electron microscopy. 10 % NBF is fine!
The overnight step in PBS is also not necessary. You can go directly into decalcification. We use EDTA here (~20 %) (between 1 and 6 days), you can also use nitric acid (6%) (maximum 3-4 days, otherwise staining of cartilage will not work properly). If you use nitric acid you then have to add a step with 5% sodium sulfate (overnight). I also think that chloric acid is a bit to aggressive; best results you will get with EDTA 15 - 20 %.
With best regards
- Malcolm Lim added an answer:3Is it still possible to do histology after inserting the catheter through heart apex?
I am supposed to measure pressure-volume relationship in a mouse TAC model
by inserting the catheter through the apical approach, is it still possible to do histology after inserting the catheter from the apex?
I concur with Frank that you can still do histology processing on the heart specimen. Fix the heart in formalin (you can mark the catheter site with a short pin or histology tissue dye which stays on after processing). As Frank said, using different color can help you mark out the orientation of the heart (inclu. the insertion spot if it matters to u).
If you are using Histology tissue dye, apply the dye, dab excess with paper towel, mist-spraying 2% acetic acid can help to "fix" the dye.Following
- Markus Merk added an answer:4What would be a good alternative to alpha-bungarotoxin for paraffin AChR at NMJ staining?
I try to stain AChR in neuromuscular junctions in paraffin embedded native mouse muscle sections and I can’t get the Ab (clone61) from Sigma and Abcam for the alpha-subunit to work. Does anybody know a good alternative or a working protocol?
Thanks a lot for your suggestion are you using mAb35 in mouse models and are you applying a special protocol?Following
- Emilio Lanna added an answer:16Does anyone have any tips on sectioning formalin-fixed paraffin embedded tendon sections?I tried sectioning cold and room temperature blocks. The tissue seems quite brittle and even with a sharp blade, I can't make a nice 5um section.
Fortunately, this technique also works with elastic marine sponges (Porifera).
- Wincenty Kilarski added an answer:5Can any histological resercher help me with the scoring of artemia tissue with H&E?
I'm using H&E.
First, please define what do you want to score? and on which part of tissue of Artemia? Hear you soon.Following
- Rozlin Abd Rahman added an answer:9Has anyone experienced negative staining on Safranin O with BMSC?
Does anyone ever come across any paper that mentioned on negative staining for safranin O for BMSC?
I am having trouble in trying to justify my result.
I am using BMSC seeded on PLGA/Fibrin scaffold to promote cartilage restoration. Other parameters (MTT assay, sGAG content and RT-PCR for collagen type II, aggrecan, sox 9, collagen type IX ) were somewhat favourable. However it was not supported by the safranin O as there weren't any orange-red staining (in vitro and in vivo). But my Alcian blue did show some cartilage ECM production.
How do I explain this event?
Thank you very much for your kind assistance, and I am very much grateful for your feedback! In fact, I too almost forgot about this question, until I got a notification from you. and for that I thank you again.Following
The study of the microscopic anatomy of cells and tissues of plants and animals, including tissue fixing, fixation and staining.