- Claire Wood added an answer:2Is 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?
Thank you! That's very useful info.Following
- Bartosz W. Schramm added an answer:7Is there an open source automated image segmentation tool for histological cross-sections?I am looking for an automated method of image analysis that would aid us in a morphological analysis of histological cross-sections of Beetle tissues. The most promising approach that I managed to find was an AdaBoost and GVF Snake algorithm, but I would need some help with getting through the mathematical formulas to a software. Any advice?
It's been almost 2 years since I faced the problem and to date I was able to solve some with ImageJ and Matlab but I will most definitely check it out. Thanks for the heads-up John. I highly appreciate it.Following
- Naziha Mansuri added an answer:2Can any histological resercher help me with the scoring of artemia tissue with H&E?
I'm using H&E.
Dear Najla, i will add that scoring methods different from subject to subject, so first you have to chose what do you want to score, then you will chose the method of scoring that you will follow, different methods yields to different results. I am suggesting that H&E used to make the tissue more clear under the microscope, specially cytoplasm and nucleus.. but scoring using H&E , THIS IS SOME THING SPECIAL AND NEW FOR ME?
hop that i can help you more.
- Sowmya Ramesh added an answer:18Does anyone have a good protocol to reduce/remove autofluorescence in tissues?I need to performe some immunofluorescent assays in embryonic mouse lung paraffin sections, fixed in formalin. However, the autofluorescent signal (both green and red channels) in the negative control is a bit high, so I cannot really validate my results. How can I solve this problem?
I was performing TUNEL assay on rabbit growth plate sections. Tissues were fixed in 10% buffered formalin followed by formic acid decalcification. For the TUNEL, I followed the recommended protocol of permeabilization with proteinase k and further labelling with the reagent and the enzyme (Cy3 for detection). After that I used the mouting medium which has DAPI in it and covered the slides.
When I observed under the microscope with DAPI filter, I find only black screen, with Cy3 filter it shows very few positive cells for TUNEL. But then with FITC as well other filters, the growth plate shows autofluorescence. I wonder if it is because of the fixative used?
To check if DAPI was the problem, I just added DAPI to growth plate tissue and to another slide where the tissue was fixed with formaldehyde. Growth plate tissue showed very very weak positive cells while the other section showed good staining.
I am not sure how to get rid of this problem.Following
- Gudrun Lang added an answer:2Does residual formalin affect my proteins stained by coomassie stain?
My whole cell lysate samples have residual formalin (from formalin inactivation for hours to days). When equal amount (based on OD600) were loaded on SDS-PAGE gel, I saw much less proteins on formalin treated samples than untreated samples. Why?
I am no expert in this field. But I know, that the isoelectrical point of tissue-proteins is shift after formalin fixation. As far as I remember the IP is lower afterwards.
Regarding the length of the formaldehyde incubation first hydroxymethylen-adducts at prefered aminogroups develope and then cross-linkings via methylen-bridges are built.
So I think you deal with rather changed proteins after formaldeyhde-treatment.
Coomassie blue is an acid dye that binds to aminogroups of proteins. These are grossly the same binding-sites as formaldehyde prefers.Following
- Joanna Stanicka added an answer:7Where is signal this from?I performed immunofluorescence staining and got the following result.
I don't know location of this signal, is it ER, mitochondria or secretory vesicle?
To confirm this, we would have to purchase each marker or tracking dye, but this is too expensive.
I'd like to get some comment or hint from anybody who has many experiences about the immunofluorescence signal.
It looks like mitochondria to me. The secretory vesicles would be a lot smaller and the endoplasmic reticulum staining looks like a a very thin network.Following
- Vyacheslav Lyashenko added an answer:19Where can I find a database of images of cytological and histological preparations?
I'm interested in the images of cytological and histological preparations. Where can I find a database of images of cytological and histological preparations?
Thank you, dear Rachel.Following
- Yi-hsieng Samuel Wu added an answer:2Could 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 agree to MahdyFollowing
- Kristine Atkinson added an answer:11Is it possible to use frozen tissue (-70C) for pathological study?
I have frozen rat kidney and I want to do a histological or pathological study. Can I use frozen tissue or I have to defrost them? If I defrost them, will the integrity of the tissue be destroyed after defrosting or not? Thanks for your attention
The structure disintegration happens when the ice melts at about zero degrees, yet many chemical activities can occur below freezing temperatures. I would try immersing some of the tissue sample in fixative at -20 degrees (or maybe minus 10, usual frig freezer temp) then go through the normal wash and dehydration series in the freezer for embedment in Lowicryl. This resin is polymerized in the freezer with ultraviolet light. Then once embedded you can do all kinds of wonderful things like immunogold and TEM.
See J Histochem Cytochem. 1984 Nov;32(11):1217-23. Rapid embedding of tissues in Lowicryl K4M for immunoelectron microscopy. Altman LG, Schneider BG, Papermaster DS
The abstract supplies adequate instructions. http://www.ncbi.nlm.nih.gov/pubmed/6436366Following
- Vyacheslav Lyashenko added an answer:2What tasks need to be addressed in image processing of cytological and histological preparations?
What are the main tasks and problems in image processing cytological and histological preparations.
How can help medical professionals image processing cytological and histological preparations.
I'm interested in the publication on this topic.
Thanks for the detailed answer.
This is a very interesting and useful for my work.Following
- Hamny Hamny added an answer:4Ho do I fix and embed ovaries?
I am looking for a protocol for fixing and embedding cat ovaries using 10% neutral-buffered formalin for use in IHC/ISH.
Ideally, I would like specifics on how ovaries are prepared (whole or halves or slices?), the length of fixation, and the temperature at which you do fixation. I am having trouble building a protocol from published methods alone.
I had the experience of IHC and HE staining. For IHC, I often use 4% paraformaldehyde fixative (for 7 days) depending on the organ. Sometime only 4 days. For brain and intestine, i conducted perfusion process with the same fixative through the heart. I have tried IHC staining with Bouin solution and 10% BNF (buffered neutral formalin) as the fixative solution and the result is good. For HE staining I mostly use Bouin and 10% BNF as a fixative solution
Good luck KathrynFollowing
- Syed Zahid Ali Shah added an answer:5Can I cool slides in refrigerator after heat induced antigen retrieval?
it takes long time till the slides reach to room temperature after heat induced antigen retrieval so can i cool them down in refrigerator?
thanks for your nice replies....if i let it cool at room temperature and then wash in ice cold buffer is that ok? and if i let it cool for bit long what will be the effect? anything can go wrong?Following
- Gudrun Lang added an answer:4In Situ Hybridization - How can I keep the larval section attached to slides?
I am having trouble keeping FFPE larval sections fixed to the slides during ISH. Most detach during the heat pretreatment and the protease digestion. Some are breaking apart, and others are coming off in one piece.
I have tried several different fixation protocols and slides. I suspect that the fatty nature of the larvae is what is making adhesion so troublesome. Does anyone have any ideas for me? Thanks.
We bake our paraffin-sections for ISH overnight at 60°C. This either increases the sticking but also the quality of ISH-result. It is some kind of "aging", that is known in cellpreparation for genetics.
For gentle permeabilisation I decrease temperature and prolong incubation time. For FFPE-sections this is usually 60 min, 80°C in citratbuffer pH 6. So we don't cook our slides.Following
- Jeanette Saskowski added an answer:14How do I get even sections with my Vibratome 1500?
Our lab has a Vibratome 1500 that was working well until a recent cross-country move. I've tried adjusting the speed and amplitude of the blade, as well as the blade angle, but the sections are always missing a portion. If you look at the blade in the blade holder, it appears that the left side of the blade is just slightly higher than the right side. And, when you start sectioning, the liquid in the specimen tray is at a higher level on the right side of the blade than on the left side. Is there something I'm doing wrong or do I need a new blade holder? If I need a new blade holder, does anyone know where I can buy one? The manufacturer is out of business and I can't seem to find anyone who services them or sells parts. Any help would be appreciated.
Thanks for the purchasing information. I've ordered a box of 250 Personna blades. Still working on the two Vibratomes....
- Taben M Hale added an answer:15Does 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.
I am so sorry for taking so long to respond...I don't sign into Research Gate very often. What ultimately worked best for me was soaking the block face down in Downy fabric softener (5ml Downy in 100ml distilled water). I got this tip from the attached document. Best of luck!Following
- John Brian Jones added an answer:11Has anyone used glyoxal as a substitute for formalin when fixing invertebrate tissues?
Glyoxal is a very old formalin substitute, it has been successfully used in medical histology (mammals), but I am unaware of papers comparing its performance with invertebrates.
Thank you for the references, they are useful additions to the debate. Its becoming clear that when we have the time :-( we will have to do our own comparison experiments.Following
- Khedidja Tair Abbaci added an answer:4Which is the most efficient protocol for processing histologic samples of xenogenic bone block for decalcified histological analysis?
Which is the most efficient protocol for processing histologic samples of xenogenic bone block (Equine Espongeous) for decalcified histological analysis? EDTA takes far way too long and Nitric acid in higher concentrations than 5% can damage the samples. Thanks in advance. Best regards.
we use 10% nitric acid and then checked by pricking it with a needle to check the hardness of the sample
- 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.
- Jennifer H Steel added an answer:7Does 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!
I would recommend not baking the slides, because this can damage the antigens. However, if you use poly-L-lysine or Superfrost coated slides it's not necessary, as long as the slides are dried at 37-40 degrees. I use this procedure for TMA and no sections come off.Following
- Manal Tawfik Hussein Mohamed added an answer:4Any 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?
I also prefer neutral buffer formalin for fixation. you should take care and adjust proper time for fixation in Bouin or any fixative, the embedding time should be also adjusted according to your samples(size, age, etc)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
The study of the microscopic anatomy of cells and tissues of plants and animals, including tissue fixing, fixation and staining.