- Yelena G Golubeva added an answer:Why are my spinal cord sections curled after IHCs?
So I'm working with mouse spinal cords and sometimes the sections are curled/rolled after being sectioned/IHC. It doesn't happen to every batch processed but a good portion do. What would cause that? The tight curling makes it extremely difficult to slide mount. Is it the perfusion? sectioning? iHC protocol? I don't know where to begin troubleshooting.
Here is my current protocol:
The animals are perfused and fixed using 4% para, the lumbar region is removed and placed in a vial of 4% para for one hour. Athens cords are then transferred to a 30% sucrose solution for 3 days then sectioned on a Lecia cryostat at 40micrometers. Sections are placed in cryoprotectant for short term storage for IHCs.
Any assistance would be much appreciated.
First aid is an antiroll plate. If it is adjusted correctly (I do it on OCT (tissue-Tek freezing media) block without tissue which you can keep at-70 for continuous use), it provides for a flat section. You can also warm up the lower cutting edge of the tissue with your finger just before making a section. Try -16 C; it works better for brain tissue. The other approach is embedding in OCT media in a cryomold leaving enough media around the tissue, cut a section and mount on a glass ++ (charged) slide. If the edge of the section curls in this case, it doesn't affect the tissue when you mounting the tissue on a glass slide. If you can use disposable knives (they are not expensive) it will help you as well.
- Musfira Uvaize added an answer:Does anyone have experience with Emerin staining in HEK293 cells?
Hi, I am doing emerin staining using HEK293 cells. The problem is that the cells are showing not only the "ring" shape of nuclei, but also have some tail-like staining around. Does anyone know what the problem is? The antibody works well in human primary fibroblasts in my lab, so it's not the antibody problem I think.
I am wondering if there is anything inappropriate in my fixation step:
HEK293 cells were seeded on the Poly-L-Lysine coated coverslips/dish. I tried methanol (-20°C 7min) and 4% PFA+0.2%TritonX-100 (10 min, RT) to fix the cells, but they all show the same staining like I said above. Any suggestions? Thanks!
Then it is probably the ER you are seeing.Following
- Sylvia Ortega-Martinez added an answer:Does anyone have a protocol to dissect locus coeruleus, ventral tegmental area and amygdala from adult mice?
I would like to dissect these brain places from adult mice but I can't find any protocol or suggestions referred to dissections.
Thanks again Mattew!
Of course it helps! As i saw in your profile that you also work with prefrontal cortex, can you suggest me some marker for this brain place?
- Stephanie Lesage asked a question:Does no sign of chronic inflammatory reaction at histology of explanted samples mean no pyrogenecity too?
In a subcutaneous implantation assay (rabbits for example), if there are no signs of inflammatory reaction upon explantation and histology, does it mean that the material is non pyrogenic as well ? What are the other minor detectable signs of pyrogenecity, other than fever ? Thanks!Following
- Anyone has experience with ThioflavinT stainign in Frozen setions?
I am trying to stain tangles in human brain frozen sections. Thioflavin T has been widely used by the people in paraffin sections, however in frozen section I am seeing lots of non specific binding. The image lights up like Christmas. Any suggestion would be appreciated.
I tried 0.5% and 0.1 % Thioflavin in 0.1N HCl.
Dear Mitesh, you're welcome.
Just to offer: if you can't access the above cited articles, let me know....
I'll provide later on 2 recipes out of Bancroft /GAMBLE,6th Ed(2008/2011 (originally applied to fixed / paraffin sections, Thioflavine T (but only, if you reply that you don't have these recipes = VASSAR & CULLING 1959 and BURNS et al, 1967= ph1.4 Th-T)
and - additionally - you should have perhaps a view into:
GUNTERN R, et al,1992_Experientia 48,p.8-10: < An improved Thioflavine S method for staining neurofibrillary tangles and senile plaques in Alzheimer's Disease > (which has been described for application to cryostat sections).
This article one can find @ http://link.springer.com/article/10.1007%2FBF01923594
(only PPV on first sight, but if you click: <Look inside>(you find the icon above the <Title> and <Experientia> you are able to see the first two pages which include
i) the method as well as ii images....).
I've found other possibilities for staining tangles, perhaps more specific than Th-S (which has been said to be +/- unspecific) like X-34 (X-34, a lipophilic, highly fluorescent derivative of Congo red), cf.:
STYREN et al, < X-34, A Fluorescent Derivative of Congo Red: A Novel Histochemical Stain for Alzheimer's Disease Pathology >
doi: 10.1177/002215540004800906 J Histochem Cytochem September 2000 vol. 48 no. 9 1223-1232, to be found as pdf free of charge/free access @:
Best wishes, good luck and regards, WolfgangFollowing
- Virginie Mauro added an answer:How should one prepare formalin solution for formalin test in mice?
Low concentration (e.g. 2% to 5%) of formalin solution is widely used for well-established formalin pain test in rodents. But can I use the same as the formalin solution for fixative (such as the 4% PFA in PBS buffer)? How do you prepare it for formalin test? Thanks a lot!
Use 37% commercial solution (buffered as possible) and make your dilution in PBS 10mM pH=7,4 to avoid pH changes due to formalin polymerization.
I used it for fixation in rodent, it worked very well.
- Is anyone familiar with cardiac stains for hypertrophy and fibrosis?
What stain can I use on cross-sections of mouse heart to 1) provide gross morphology 2) Determine cell size of individual myocytes (myocyte area) and 3) estimate changes in fibrosis. Will Trichrome work for all 3 purposes?
Dear Chad, dear Aleksandar,
the article cited above by Aleksandar is really nice and shows some excellent images of sections processed as described.... Chad, perhaps you have gotten meanwhile the / a pdf for personal use by Aleksandar. For all others just for their convenience: the Article's Abstract: can be found at: http://informahealthcare.com/doi/abs/10.3109/10520295.2010.528026,
unfortunately BT&H (former title: Stain Technology)-articles are only accessible either by subscription or PPV (pay per view).
Have a nice day and a successful week, best regards, WolfgangFollowing
- Fateme Parvizi added an answer:Can anyone help me about histology of mantle in pearl oyster?
I need some information about mantle cells.
Im very appreciate
I have these papers except second one, but these have not adequate information for my project.
thanks for your helpFollowing
- Abdelraheim Attaai added an answer:Does anyone what is the best cutting Thickness for microtome?
i want cutting the gill and liver tissue of fish
I agrre with all above answer. I want to add that if you are a beginner, you should better start at thicker sections (about 7-10 µm) and gradually shift to thinner sections till reach 3-4 which is preffered for traditional stainings for light microscope. thicker sections will give you better images for low magnification and thinner sections for higher magnification for detailed cell analysis.
- Cheryl Seifert added an answer:Why does an H&E stained slide that showed the presence of calcium does not stain positive for von Kossa?
An arterial plaque was partially decalcified; one section was submitted for H&E stain.H&E stained slide showed the presence of calcium in the tissue; however, when stained with von Kossa; the result was negative. Then the very same slide was re-stained with H&E and once again the slide showed the presence of calcium. Why is it so?
This technique is for demonstrating depostis of calcium or calcium salt so it is not specific for the calcium itself. In this method, tissue sections are treated with a silver solution and the silver is deposited by replacing the calcium reduced by a strong light, and thereby visualized as metalic silver.
1)If stain is weak or possibly rinsed off during the washing steps, it is indicated the UV light wasn't strong enough.
2)Oxalate salts are usually believed to give a negative von Kossa staining.
3) A negative control may be needed when there is any doubt that the resulting black deposits are calcium. The is done by treating a test slide in 10% formid acid for 10 minutes prior to incubating in silver stain. The test should show a negative reaction.Following
- Marina Yurieva added an answer:Does anyone have an idea what can be the problem during our RNA ISH on free-floating mouse brain sections?
We do RNA ISH on formalin fixed mouse brain slices (not paraffin embedded or frozen)(40 µm) with DIG-labelled RNA riboprobes as described in the attached file.
The problem is that the success of the protocol is about 40%. Once we got sign, then there is no signal, even if the circumstances are almost the same. May be the duration of perfusion with DEPC treated PB(± 2 min) and PFA-fixed brain slicing differ time to time, could it cause significant mRNA degradation in the tissue?
The other possibility we thought is the degradation of the probes. We store them at -20°C and dilute during ISH as described in the protocol (in the attached file). What is the best way to check the integrity and functionality of the probes? Or could be something else the problem?
We started using this technique recently, so we have not so much experience in RNA ISH. We appreciate every help!
If it's 4% PFA overnight fixation, it should be at 4°C or only for a few hours at room T. I was working on whole 9.5 dpc mouse embryos and was transferring them to methanol for longer storage but if you just fix them with 4% PFA staining is usually better.
Probe itself plays a big role: some probes work right away and don't have any problems while you have to incubate others with NBT-BCIP longer. You can try it with a different probe, longer incubation time and higher antibody concentrations. Basically you can try and tweak every step of the protocol, for me it was always much easier to make a new probe than to spend a lot of time on checking every parameter.Following
- Joao Paulo Rodrigues Marques added an answer:How I can prepare slides for the study of plant histology?
How can I prepare slides for the study of plant histology? Please also suggest the other precautions and reagents required for this study.
There are several ways to prepare slides for hiistological analysis. I notice that you want to detect physiological changes, so you will need to embbed your samples in resin ou parafin.
I will give you a standard method for light microscopy.
First of all, you will need to colected the samples and fixate as soon as possible. You can fixate in several reagents. I'll give some examples: you can use FAA (Formaldeid, acetic acid and Alcohol 70% or 50%, or youn can use Karnovsky solution ( Karnovsky, M.J. 1965. A formaldehyde-glutaraldehyde fixative of high osmolality for use in electron microscopy. J of Cell Biol 27, 137-138. )
During the fixation the samples needs to be submitted to a vacuum pump to remove the air inside the samples. I use 15 minutes for 3-4 times during the day.
After that you made the dehydration in an ethanol series (10, 30, 50, 70, 90, and 100%). - 10- 20 minutes in each ethanol grade.
When you finish the dehydration you need to make a transition of ethanol to resin (3:1, 1:1, 1:3,) the time for each step is variable. Depends the size of your sample. you can write to me indicating what plant tissue do you want to analyse. What is the size? I use a plastic resin (Leica Historesin®, Heraeus Kulzer, Hanau, Germany). But there is other companies that produce methacrylate resins.
But if you will use paraffin, you need to make a transition of ethanol to xilol (3:1, 1:1, 1:3) (one hour per step) and than other transion for xilol to paraffin (3:1, 1:1, 1:3). When you use paraffin you need to have one Oven at 60 C. Because paraffin is solid at room temperature. I send you a good protocol about paraffin in attached file.
The polimerization of resins happens at room temperature.
For cut your samples to do the slides you need a microtome.
Please let me know if you understand each step. If you have some questions feel free to contact me.
- Taras Kotyk added an answer:Is there any literature outlining the measurement of aorta diameter when the shapes are not fine round?
I want to measure aorta diameter but the shape is not find round.
Found geometrical centr of lumen and calculate with macro distance from this center to all points at the lumen selection, mean of this distances will be radiusFollowing
- Taras Kotyk added an answer:Measuring adipocyte cell size in a suspension from pictures - can anyone help?I want to use automated programs such as imageJ to quickly measure adipocyte size. The adipose tissue has been digested with collagenase and the adipocytes are floating in saline for the microscopy pictures. Look at the attachment for an example picture. Is there an existing macro/program to quickly analyse the diameter and/or are of these cells? Most programs for imageJ (say http://sw.wikkii.com/wiki/Adiposoft) are used for stained histological sections. The adiposoft macro does not work for pictures in suspension that I want to analyse.
How should I proceed or is there a ready made program for this?
Try such algorithm, modify, create macrotoolset...
1. Split Channels
2. Select red channels
3. make binary
4/ binry option -> dilate
5. Fill holes & manual correction
6. remove outliers
8. analyze particles (circ 0.75-1.00)Following
- Gloria E Hoffman added an answer:Can ice-damage be completely avoided when embedding frozen brain tissue and how?
Recently we are desperate about ice-injury of mouse brain slice in HE staining. There are so many small holes in our 8um slice, probably caused by ice-formation. We perfused our mice and used 20%(mass ratio, solute/solvent) sucrose to dehydrate their brains, still a lot of holes as a result. At freezing step, dehydrated brains were transferred immediately into -80 centigrade fridge, maybe it was not quick enough?
But some published paper also display photo with similar injury, so can this be fully avoided and how to mitigate as much as possible?
Thanks in advance for any advice!
We have used 30% aqueous sucrose which prevents the ice crystal formation if the tissue has been in the sucrose long enough (a few days usually for brain) it must sink or some weight put on it to keep it immersed. Freezing can be effected on a dry ice copper stage or the other methods mentioned above.Following
- Olga M Pulido added an answer:Where can I buy a decent set of prepared microscopic slides with human tissues?
I'm trying to instruct students (informally) on using a microscope/basic histological techniques/basic identification of disease states in humans.
I've found quite a few prepared microscopic slides with human tissue, but none of them seem to come with a book or informational material to help the students understand what it is they are looking at/what makes a certain tissue diseased, etc. Will I have to purchase a separate book and the slides or is there a comprehensive set that would work for this purpose? Ideally, I'd like human tissues along with diseased/non-diseased tissues for easy comparison.
At the time of digital pathology and for educational aim is matter to explore universities with large pathology departments both human and veterinary schools. Long ago before it was closed the Armed Forces Institute of Pathology US had the best collection of histology slide including the text and publications. May be some one knows what happened with that material or if there is any one else managing that or similar collection. I personally sill prefer to train students with slides and microscope on hands but digital material could be a reasonable alternative.Following
- Abdelraheim Attaai added an answer:What is the best orientation to cut sections of mouse intestine?
Does anyone have a preferred orientation in which to section mouse small intestine so that one maximizes the number of intact villi in the section? I have tried paraffin-embedding the entire "tube" and cutting transverse sections but with little success. I have now fixed sections where I have cut the tube open such that the villi should all be on one face of the section and the mucosae muscularis on the opposite face. What is the best way to orient this type of section in paraffin to see the most villi? Thank you in advance for your help.
I agree with Azza and Sebastian provides a good idea.Following
- Saleh Alkarim added an answer:Is anyone familiar with heating a Corning 24-well tissue culture plate?
Does anyone know if a Corning 24-well tissue culture plate can be heated to 37-degrees C (98.6-degrees F) without any issues? The last thing I want is the plate to melt, or some other catastrophic event to occur, while trying to incubate tissue in 2N HCl....
i agree with the researchers , Corning 24-well can withstand temperatures higher than 37 C with 2N HCl
see the attachment
- Amit Dubey added an answer:Can anybody suggest the best method for oil red-o staining of differentiated adipocyte cells derived from Wharton's jelly stem cells?
I used thermoscientific protocol for stining but it did not give result well , can anybody suggest some different protocols for staining.
Thank You Mahmoud........Following
- Chee Won Oh added an answer:Is there any Histologist out there with expertise in skin?
I am looking for someone who has experience in examining skin samples post mortem. In particular structural changes or differences between samples so a histologist perhaps?. If anyone has the skills and knowledge to do this or could point me in the direction of someone that does then that would be great - Thanks
- Vitus Stachniss added an answer:Why after MMA embedding some samples are completely polymerized and others are rubbery or not polymerized at all?
To embed vertebrae, long bones and calvaria I fix in PFA 4%, dehydrate and after double infiltration in a solution of Methyl Methacrylate (MMA destabilized+NPG+BPO), I use the embedding solution consist of MMA destabilized, NPG, BPO and DMPT. Sometimes happens that among the samples simultaneously embedded just some solidify and others have problems. Can anyone help me to understand how obtain perfect embedded samples?
We observed similar problems with Kulzer MMA Technovit®9200 when embedding hard and soft tissues. We could improve resin polymerization with a pressurizing device (6,5 kg/cm^2) as described in: A: Steiniger et. al. Ti: Immunostaining of pulpal nerve …. embedded in Technovit 9200: Cells Tissues Organs 2013;198: 57-65 and www.karger.com/doi/10.1159/000351608
- Does anyone have a copy of the instruction manual for a Lynx EL tissue processor?
I have just bought a second hand one, but with no manual.
Dear Alan (dear Ranu, (:-)) ),
sometimes it might be better to wait a bit for a reply....if one knows more in detail about the problem perhaps <help> is easier than expected...
hoping I might help you really in this case...see attached file:
LYNX Service/operating Manual as of Sept.1996, by courtesy of an excellent REICHERT-LEICA Service Rep (unfortunately retired for years now....).
Best wishes & good luck, WolfgangFollowing
- Kieran Mcallister added an answer:Why 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 used to do a lot of resin work and this kind of problem was usually caused by the final consistency of the polymerised resin. You seem to have a good protocol, so assuming that you are not having sectioning issues, the only possibility that I can think of is that your resin is too hard or inflexible. I know a retired gentleman in the UK who is an absolute genius with resin histology. His name is Neil Hand, formerly of Nottingham Hospital, but he is still an assessor for UKNEQAS ICC and may be contacted through them I'm sure.
Best of luck,
- Raghuveer C V added an answer:Can anyone help with a problem in histology paraffin tissue sectioning?
Hi.. I have encounter a problem in histology tissue sectioning.. tissue processing is good and section also is coming properly but the problem is after drying the section alone coming out from the slide but the paraffin is fixed properly. what might be problem in this case?
Go through every step of processing carefully. Make sure about temperatures. Select the right type of paraffin wax with ceresinFollowing
- Melissa Laroche added an answer:Does anyone have success with Anti-FLAG antibodies in brain sections?
We have mice expressing a Flag-tagged protein (only one flag epitope) at endogenous levels. We have major problems (non specific labeling; particularly strong in the olfactory bulb) with most antibodies in our perfusion-fixed brain sections. Anyone has ever used anti-FLAG antibodies to label mouse brain proteins? Any advice? Best fixative to be used? Aware of a specfiic antibody?
As discussed with one of your student today. Despite the blast made with your flag in the olfactory bulb, there still appearance of non-specificity. Maybe to testing with blocking solution more concentrated. Currently, I understand that you work with the 3% but often with my olfactory bulbs blocking solution is between 5 and 10% depending on the case.Following
- Yuefeng Tang added an answer:Is anyone familiar with Cre/loxP for gene conditioning knockout mice?
We are creating a gene conditioning knockout mice strain by Cre/loxP. I think the first step is to examine the physlogical difference between it with WT. What parameters should I examine? such as weight, food intake, vital organ histology? anything else?
Thanks so much.
If the conventional knockout mice are lethal or if not, I'd like to use CMV-Cre to delete the gene to test the function (if lethal) and confirm gene deletion.Following
- Iker Rodriguez-Arabaolaza added an answer:Does anyone know of any Anti-Pig Endothelial Abs?Does anyone know Endothelial antibodies (like CD31, Von Willebrand, VEGFR or VE-Cadherin) that work well in pigs?
Thank you very much in adavance for your help
Thank you very much to all for your answers. I really appreciate the information.
- Fang Li added an answer:Why do we still need to use salines after fixing in histochemistries?
After the fixation with 4% paraformaldehyde (PFA), I assume that everything that is fixable by PFA will be preserved while those did not will lost much of the attention from the investigator (may due to degradation or assuming lost of physiological characters).
Interestingly, according to what I read/experienced so far, people usually use saline or its associated form as the main buffer solution during the histological staining procedure, particularly for the immunostaining parts.
I was wondering what would be the main purpose/advantage of using saline at the post-fix stage. Additionally, I was wondering whether people had tried using ddH2O instead of saline in the steps which originally would use saline.
Thank you very much in advance!
I agree with Francisco. Even after fix, the antibody and antigen and their reaction also need proper buffer to maintain the condition. We generally use PBS.Following
The study of the microscopic anatomy of cells and tissues of plants and animals, including tissue fixing, fixation and staining.