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Questions related to OCT
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We are looking for a copy of the OCT Reader java plugin for ImageJ that
was distributed by Bioptigen. Does anyone have a copy they could give us?
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Hello,
I am trying to extract high quality RNA from fresh frozen OCT samples/sections.
These are small cartilage samples and succeeded only once (out of what seems to be 1 million attempts). I've trimmed off as much OCT as possible, and even picked the sample out of the OCT when sectioning.
My 260/280 ratios aren't that bad ~1.8, but the 260/230 ratios are consistently poor and around 0.5.
Do you have any suggestions on what this could be and/or how to improve the quality?
Below are the things that I've tried
- Trizol with and without colums
- Biomashing or bead homogenizing
- Chloroform extraction
- 15um sections, 25 sections into 350uL or 1000uL Trizol
Thank you!
-Merissa
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I think it would be better if you extract the RNA without embedding in OCT (if you work with the whole sample, not with sections).
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Hi,
I OCT embedded some cerebral organoids but am having issues sectioning them. Is there any way to take them out of the OCT and do a paraffin embedding to get thinner sections that are more well supported and give better structure?
Thank you
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I think you can just unfreeze the OCT block and take the organoid into ethanol for some days, since you have to dehydrate it before putting in paraffin. Before paraffin, you have also a step in butanol (ON) or xylol (15 minutes) to make the tissue harder. Then, you can put in paraffin for some days (it can stay for long time without problem). Finally, you can do the paraffin block.
I hope it works for you.
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1. The necessity of a polarization controller for single-mode fiber. Is a polarization controller necessary for single-mode fibers? What happens when you don't have a polarization controller?
2. Optical path matching problem. How to ensure that the two arms of the optical path difference match, any tips in the adjustment process? If the optical path difference exceeds the imaging distance, will interference fringes fail to appear?
Only these questions for the time being, if there are more welcome to point out.
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1. The polarization can change during propagation. This will degrade the visibility of the fringes. You need a polarization controller or a polarization-maintaining fiber.
2. By the imaging distance, do you mean the distance to the sample or the sample's dimension? In any case, if the OPD exceeds the coherence lenght of your source, the interference fringes disappear. In order to match the OPD, we typically sweep the reference arm a long distance and record the output intensity. Another method is to monitor the output spectrum using a spectrometer. The spectrum shows oscillations for non-zero OPD. When the OPD approaches zero, the spectrum oscillations tend to disappear.
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I have a consistent issue of having folds in my cryosections for some of my mouse breast tumour samples.
Sample processing up to this point is:
-2 H 4% PFA fixation at RT.
-2x 1H wash in DPBS at RT and O/N DPBS wash at 4C.
-30% sucrose incubation for 24H at 4C
-30% sucrose:OCT (1:1) 1H at RT
-cryoembedding in OCT.
-Stored at -80C
-1H equilibration to -20C before sectioning.
Cryochamber temperature was tested at a range of -16 to -21C.
Unfortunately the specimen head thermostat can’t be changed and the display is broken, but I think it’s between -15 to -20C.
These sections were cut at 6um using the brush method as I find that using the anti-roll bar causes too many tears in these sections.
hoping the collective ResearchGate mind has any suggestions :D
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So I can answer my own question! In the end what worked for me was placing the microscope slides in the cryostat chamber to cool so that I can place the sections on top and gently brush the flat with a soft flat wide brush. Then with a finger on the back of the slide to melt the section onto the slide for a few sections and keep the slide in the chamber until you have enough sections.
still open to other suggestions though!
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Hi,
I would like to know if it is possible to embedd tissue (lung) which is already frozen in a cryovial in OCT for cryosectioning?
Do I need to defreez the tissue, wash it with sucrose and then freeze in OCT or is there a way to circumvent the defreezing process?
I just need to do a Oil Red O Stain, but would like to have nice stainings.
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You can add the lung tissue directly to the OCT. However, the histology from tissue prepared in such a manner is likely to be of poor quality. Ideally, the lung tissue should be inflated with OCT to expand the alveolar spaces.
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I would like to extract chromatin from frozen samples, but most of them are embedded in OCT?
Does anyone has experience with it?
Thanks a lot!
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Hélène Neyret-Kahn how did you remove the OCT from the frozen tissue to further extract chromatin. Did you use a specific kit?
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what is considered a "Good" OCT for an RFNL?
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In addition to the previous reply, here are the normal variants : The mean value for RNFL thickness in the general population is 92.9 +/- 9.4 microns. Typically, a normal, non glaucomatous eye has an RNFL thickness of 80 microns or greater. An eye with an average RNFL thickness of 70 to 79 is suspicious for glaucoma.
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I'd like to flash freeze PFA-fixed mouse brains in OCT. Usually, this is done by flash freezing with liquid nitrogen, or dry ice. I was just wondering if it would be enough to transfer the samples in OCT straight to a -80oC freezer instead? I'd like to avoid any ice crystal formation.
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You can gradually change the temperature,Initially move the tissue to PFA with 4°C then to ice chilled PFA and finally to -80°C. How ever I am not sure about crystal formation until the tissue is dry because the PFA also contains water percentage in it. If you change the temperatures without using PFA may allow the tissue to absorb moisture and sudden change in temperature may cause changes in tissue architecture which may affect the sectioning reporting. Get more suggestions form any expert pathologist.
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I have tried fixing in PFA 4% for an hour followed by 2 PBS washes, it has worked but not all the times and I end up losing valuable samples. I have seen comments that indicate to do what I´ve been doing and add an acetone step 10 mins at -20°C. I would appreciate any other recommendations.
Thanks!
Karen Garza
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Hi Karen,
I can identify two possible problems: a) 10 𝜇m is a thick section and may detach from the slide, 5 𝜇m is the usual thickness for histological examination; b) I never used liquid nitrogen because can alter the tissue properties, affecting subsequent cutting. I embed with the following steps: cold metal chunk, layer of OCT up to gel consistency, tissue section, cover with OCT, metal weight on top without pressing, freezing fluorocarbon spray on the metal weight. You should have a flat surface with tissue well integrated in the OCT. If cutting still difficult, gently adjust temperature with gloved thumb -22 might be a little too cold for certain tissues (I do not have experience with arteries) and you can try -18.
I hope it is helpful.
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The alginate hydrogel was fixed in 4% PFA for 2h, dehydrated in 30% surcose overnight, and embed in OCT. After snap frozen, the alginate hydrogel was cryo-sectioned onto a charged glass slide. However, during HE staining, the samples will detached from the slides. Is there a way to prevent the sections from detachment?
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You can try increasing the concentration of PLL used or by coating with gelatin as mentioned before by the researcher. The slides need to be socked in PBS or water to remove OCT and sucrose carefully before proceeding for staining otherwise chances of detachment are more.
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Hi everyone. I would have to cut serial sections of biopsies in the cryostat microtome to perform some immunoenzymatic methods on them. I wish I could cut some sort of "ribbon" of sections like with paraffin sections, but I can't. I tried to use the anti-roll plate but it didn't work. About 1 time out of 10 I can create a sufficiently long tape (at least 5 serial biopsies), otherwise the sections come off continuously with each other. I use the Leica CM1950 cryostat with OCT Leica FSC 22 Clear, Feather S35 blades. Thanks to those who will help me.
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I think it makes sense to use the anti-roll plate, but you have to take care that the postion of the roll plate and the kinfe edge ist optimal. To find this out you need several trials to optimize it. The speed of sections is importenat too. If you cut to fast the sections will curl, if you cut to slowly the sections will crush.
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I have a bunch of precious samples that appear underfixed for immunofluorescence with a particular antibody. They are all in frozen OCT blocks now. Should I melt them down and re-fix them or can I just post-fix the tissue sections on slides for 1 hour or longer?
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Thawing-refreezing cycles usually contribute to protein degradation and increased unspecific staining and artifacts. At this point your best option may be to cut the tissue using a cryostat, mount the tissue, post fix it and stain on slides. For unfixed frozen tissue, post-fixation might need some troubleshooting, since different epitopes are more easily fixed. A mixture of 50% methanol/acetone at -20 C for 20 min works for some samples ( https://www.cellsignal.com/learn-and-support/protocols/protocol-3589-if) or for 15 min RT with 4% formaldehyde (https://www.cellsignal.com/learn-and-support/protocols/protocol-if) If your protocol requires the proteins to be crosslinked. Remember that prolonged fixation might mask the epitopes and thus you might need to perform an antigen retrieval.
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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? 
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يمكنك مراجعة المختصين في هذا المجال
شكرا لكم
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Dear colleagues,
I am currently looking for used but minimally working microscopy, histology and cytology equipment. Is there anyone who wishes to get rid of old things? E.g. microtomes, tissue processors, microscopes, etc?
Thanks.
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من البكتريا
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We are going to use the Heidelberg OCT Spectralis for mouse OCT acquisition. However, we find several papers reporting the use of this instrument for rodents, we are not able to successfully do it. We have two separate devices (1- Heidelberg OCT Spectralis and 2- Heidelberg Hra 2). The Heidelberg OCT Spectralis has two lenses one 30° Standard Objective Lens and an Anterior Segment lens. The Heidelberg Hra 2 has 55° widefield lens. unfortunately, the 55° lens could not be assembled to Heidelberg OCT Spectralis.
As far as we know, given the high dioptre of the mouse eye, we should use the 55° widefield lens. However, using the standard 30° we get a rather acceptable cSLO image, no OCT image is displayed.
Can anyone help solve this problem? We already tried using an additional lens in front of the device lens but still not working, however, maybe the total dioptre of the lens was not enough.
Also, a paper suggests minor software modifications (using Alt+Ctrl+Shift+O in Heidelberg Eye Explorer software) which we could not figure out how that should be done. (Spectral domain optical coherence tomography in mouse models of retinal degeneration. Invest Ophthalmol Vis Sci. 2009 Dec;50(12):5888-95. doi: 10.1167/iovs.09-3724.)
These are some papers about using the Heidelberg OCT Spectralis for rodents:
1- Quantitative Analysis of Mouse Retinal Layers Using Automated Segmentation of Spectral Domain Optical Coherence Tomography Images. Trans. Vis. Sci. Tech. 2015;4(4):9. doi: https://doi.org/10.1167/tvst.4.4.9.
2- Tracking Longitudinal Retinal Changes in Experimental Ocular Hypertension Using the cSLO and Spectral Domain-OCT. Invest. Ophthalmol. Vis. Sci. 2010;51(12):6504-6513. doi: https://doi.org/10.1167/iovs.10-5551.
3- Giannakaki-Zimmermann H, Kokona D, Wolf S, Ebneter A, Zinkernagel MS. Optical Coherence Tomography Angiography in Mice: Comparison with Confocal Scanning Laser Microscopy and Fluorescein Angiography. Transl Vis Sci Technol. 2016 Aug 18;5(4):11. doi: 10.1167/tvst.5.4.11. PMID: 27570710; PMCID: PMC4997887.
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Thank you for sharing the videos. However, I do not get my answer yet. Do you look at the papers I mentioned in the question?
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I am looking for a good RNA isolation method. Most methods I have found take the whole tissue section to make RNA. However, I would like to take a small region of interest from a tissue section to make RNA.
I first dissect out my tissue, then freeze using isopropanol and dry ice. I embed in OCT, then store in -80. From there I am able to take 100 um sections to find specific sections of interest. But I need help dissecting/isolating a small portion of this section for consistent and good quality RNA extraction.
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Like Jonathan said you can just cut the ROI with a blade, as it seems to be large enough to be visualised by eye (or stereomicroscope). Or else if you have access to a laser-capture microdissection apparatus you can do it that way, once your sections are taken.
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I am sectioning brain tissue. During this process sometimes the tissue separates from the OCT or there will be bubbles in brain tissue folds that make the section wrinkle more. These can hinder the integrity of the tissue I want to analyze with immunohistochemistry. When preparing tissues to section, what are the most important details in your protocol that can help fix these issues?
Here is the protocol that I use:
Post dissection Preparation
  1. Transfer brain to a 15mL conical tube containing ~10mL 4% PFA.
  2. Refrigerate brains overnight at 4C in PFA.
  3. The following day, decant PFA and replace with ~10mL 30% sucrose.
  4. Refrigerate brains overnight at 4C in sucrose.
  5. The following day, decant sucrose and replace with ~10mL 30% sucrose.
  6. Refrigerate brains in 30% sucrose until ready to embed (at least 24hrs, up to 1-2 weeks).
Embedding Procedure
  1. Decant sucrose and place brain on a kimwipe, gently pat dry to remove residual sucrose.
  2. Fill tissue block about ½ way with OCT.
  3. Using large forceps submerge brain into OCT, fill the chamber the rest of the way with OCT to cover the brain entirely.
  4. Using an Erlenmeyer flask on ice (with aluminum foil covering the opening), puncture the aluminum foil with the Cytocool nozzle and spray Cytocool into the bottom of the flask. (This will prevent any Cytocool from aerosolizing and spraying back at you.)
  5. Pour liquid Cytocool from flask into a large petri dish on ice, fill about ½ way.
  6. Place tissue block with OCT into the large petri-dish with cytocool on ice. Can add more cytocool if necessary until it reaches about ½ way up the tissue block. Allow OCT to set, will turn completely white when frozen.
  7. Wrap frozen tissue blocks with aluminum foil and label with cryo-labels.
  8. Place tissue blocks in a box and store at –80C until ready to section.
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Dear Britanie,
I agree with Amir that it helps to trim the brain down to a region of interest if possible. I have found that one step that helps my tissue remain in the OCT compound is to replace the sucrose with OCT before placing it in the mold with fresh OCT. It is not enough to simply blot the sucrose off the tissue. I do this by placing the tissue on a glass slide and then covering it with OCT, and then carefully stirring, or simply turning the tissue over, to dilute out the sucrose. If you watch this process under a dissecting microscope you will see the Schlieren lines gradually disappear, and also be able to remove the bubbles trapped in the tissue, either with the tip of the forceps or just with time.
Good luck with your sectioning! Jill
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I have recently discovered that a good chunk of my brain tissue has holes in it. So much so that it almost looks like swiss cheese. We have had this issue in the past and we concluded that it was because our -20 freezer was malfunctioning. Ever since then, i have been storing my brains it in our main lab (-20) freezer (which is temperature monitored) and its happened again! This time a student brains were affected as well. I would like to conclude its frost damaged once again and our main lab freezer is also malfunctioning however, I would like to rule out every possibility. Is there any other possible reason why my tissue could look like this?
The specific steps I took were:
1.) Perfusion with 1xPBS then 4%PFA in 1xPBS
2.)Fix heads with ferrules overnight in 4%PFA in 1xPBS -leave in 4 degree fridge
3.)Extract brains and leave overnight in 4%PFA in 1xPBS-leave in 4 degree fridge
4.)Transfer to 30% sucrose - left brains in sucrose for 1 week
5.)Freeze in OCT in tissue blocks at -20 (main lab freezer) - brains were left if freezer for 3/4 weeks
6.) Section on cryostat (-19 to -21) and store in 1xPBS + 0.02% azide in 4 degree fridge
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For whole brains or large tissues I would not freeze your tissue in the -20 or in dry ice as the freezing process will not be even and can result in holes and other tissue issues. As Subhash suggested, I would skip step 2 and go straight to removing the brain tissue and post fixing it overnight. Then switch to 30% sucrose as you are doing but only till the tissues sink to the bottom of the container. Once they sink, they are fully cryopreserved. Now you will place the brains in molds in OCT, however; for even freezing you want to use an isopentane (2-methylbutane) bath in liquid nitrogen. To see how this is done, please refer to this video.
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On en-face OCT images this difference is easy to visualize.
In the retinal vessels the intraluminar content is hyperreflective. In the choroidal vessels the intraluminar content is hyperreflective.
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You may consider two additional factors: 1) blood absorption of NIR light is higher due to thicker medium (larger caliber) of choroidal vessels; 2) interference fringes pattern (and consequently OCT signal) may vanishes due to much higher blood flow speeds within the choroidal vessels than within cappilaries within the retina. You may follow this publication doi: 10.1364/BOE.10.000050 for more info.
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Histologically, rat retinal blood vessels were reported to be running in the nerve fiber layer. Looking at OCT scans, I believe they dig deeper or at least making groove like path in inner plexiform layer (IPL, "blue line" of different lengths under the blood vessel "BV' and under the nerve fiber layer "NFL"). This is distorting the total retinal layer thickness measurements. I could find a comment in the literature. Any feedback is greatly appreciated?
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Thanks for sharing this scan. This is located in RNFL and GCL but IPL is posterior to it and differentiation of IPL became difficult because of shadowing. Perhaps this is a Retinal Vein that lie deeper than arteries
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Hi! I am planning to do some whole mount immuno for mouse embryonic tissue. I read about ScaleA2 and it seems like a good yet cheap option, particularly as it will preserve fluorescent proteins whereas I don't think alternatives like BABB will.
My first question: For the ScaleA2 protocol, I have read that I will have to incubate my samples in sucrose solution, followed by imbedding in OCT compound, freezing and thawing prior to clearing with ScaleA2. I was firstly wondering what the point of freezing is, if you're going to thaw your samples anyway? And if freezing is not necessary, then why use sucrose at all?
My second question: at what point should one put on their antibodies? I was planning to immunolabel prior to putting the samples in ScaleA2, but won't the sucrose solution I incubated my samples in impair the diffusion of the antibodies?
Any questions would be greatly appreciated!
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Ana Rolo , may I know, how do You proceed with tissue sectioning of the samples for ScaleS protocol? I cannot find full protocol from A to Z, starting from perfusion and ending at imaging of sample.
I would need to clear brain sample, but I need to have it done on 105 µm thick sections. Should I cut them before IHC and then clear, or IHC, cutting and then clearance?
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I am looking for skin images (cancer, normal skin, psoriasis, etc.) obtained with optical coherence imaging.
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My coworker are developing skin OCT images. Please feel free to contact with me if you need more data.
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Besides Hyperreflective Foci (HRF), there are some dots are known as hard exudates or lipid-laden macrophages (especially in DME) in OCT images. How can we find out which dots are just HRF?
For instance, I attached one DME image.
Thanks
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HF are well circumscribed, discrete lesions having similar reflectivity as that of rpe layer. Unlike hard exudates they can be present in any layer of retina. Overlaying a colour fundus image on the oct scan containing the HF can help you in ruling out hard exudates.
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Aside from embedding brains in an OCT block, are there other ways to section multiple brains at once?
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Jasmine Pahle, I tried it out yesterday on two extra mouse brains and it worked. I'll now try it with rat and stay mindful of sticking to two or three brains at a time. Thanks!
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Dear colleagues I am looking for a freezing tissue protocol which can preserve good morphology and antigenicity for immunofluorescence. The scientific literature is scarce on this topic. I am not sure how to snap frozen, liquid nitrogen or isopentan? How to store samples, in -20, -80 or liquid nitrogen, which option is better? OCT cryopreservative should be used to store samples? Can it improve morphology quality?
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Do you mind post one publication you used with the acetone protocol. I got very interested in try it.
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Hi,
I have always embedded my mouse brains (PFA fixed followed by a 30% sucrose bath) in cryomolds with OCT (Tissue-Tek) before freezing and cutting with a cryostat.
My colleagues however flash froze the brains in isopentane after the 30% sucrose bath.
Is it possible to cut brains without embedding? But just attaching a side to a drop of OCT? Will the sections not break and attach to the Superfrost slide?
Or is it possible to embed in OCT the frozen brains before cutting? In that case, would you just cover your sample with OCT inside the cryostat?
Thanks,
Sophie
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Yes you can embed the brains afterwards in OCT or something else like gelatin.
Some people also cut the brains directly, only using a tiny bit of OCT to fix the brain on the holder.
You can take the brain, put it in a mold (i did mines myself using aluminum foil), put oct on top and let it set on dry ice. Once it is starting to set, pop it in the -80C freezer.
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I have cryopreserved OCT mounted human nasal polyp tissue slides, which I want to study post-immuno-fluorescence staining. I have the following concerns-
Whether an antigen retrieval step is required? If so how to retrieve antigen?
Which anti-fade/mounting medium should I use?
I will appreciate it if someone could suggest a good protocol or at least addresses my concerns. Actually, the protocol, I saw, is not suggesting any antigen retrieval step. Further, most of the available anti-fade mounting media are xylene based and I guess, for OCT frozen tissue, staining does not require xylene. Therefore, I have a concern about the use of these anti-fade mounting media.
Thank you
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Hi Mohammad Asad , this is the protocol I use, for animal tissue in OCT at least. you can start by washing the tissue in phosphate buffer (physiological pH). You should also wash in this buffer between each step. Then block for 1hour in 1:100 dilution of bovine Serum albumin and normal goat serum. Wash and incubate in primary antibody for ~18 hours (overnight)(this step will need to be troubleshooted based on the antibody you are using), then incubate with secondary raised to the primary antibody, for one hour. (The secondary antibody should be biotinylated) Then incubate in avidin biotin complex. This complex needs to be pre incubated for at least 10-20 minutes before being applied to the tissue (1:500 avidin and 1:500 biotin), once the complex is formed, let it incubate on the tissue for 1h. Then, add your PE dazzle 594 conjugated to streptavidin and incubate again for an hour. Hope this helps!
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I injected my rat via intravenous tail vein injection with an AAV2-CBA-GFP virus... waited 14 days for viral infection and then perfused my rat with PBS to wash out the blood and 4% PFA to fix all tissues. Post fixed the liver overnight in 4% PFA made in 0.2M PO4, sucrose cryoprotection, embedded in OCT and sectioned 10 µm sections.
When i was performing GFP staining with anti-GFP primary antibody and Alexa-488 conjugated secondary antibody I could not see any GFP signal in my green channel. All cells were fluorescing green, even in my negative control sample where I did not inject the virus.
I used 0.3M glycine to quench during my IF protocol.
Can anyone help me in reducing background fluorescence in the liver?
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TrueVIEW Autofluorescence Quenching Kit works well for liver tissue. Good luck!
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Hello,
Like the title says, I'm staining chicken sperm storage tubules with H & E to prepare for LCM. My tissues are frozen and sectioned via the cryostat with OCT. I just tried staining for the first time using the Histogene Frozen tissue staining kit by thermofisher, and my slides have a white chalk on them. I read online that this could be that the OCT has not left the slide. Is this correct? If this is the case, can I remove it with tap water after the slide thaws and before I start the dehydration process with 75% ethanol (as per the kit's instruction/protocol).
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thank you! I will try that today! Meetu Wadhwa
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When doing IHC-F, I found a lot of ugly holes on my mouse brain slices(20um). That's really bad and I don't know how to avoid the damage to my samples. After perfusion fixation, 4% PFA were used for post-fixation for 24h, followed by 24h dehydration with 30% sucrose solution. The brains were then dried with dustless wipes and embedded with OCT, to be frozen in -80℃. Anything wrong with my steps? And how to adjust them?
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These holes are caused by the enlargement of blood vessels due to the excessive pressure of fluids used in perfusion. Formaldehyde subsequently fixes the blood vessels in this position. Reduce the pressure of fluids by using a thinner needle for perfusion.
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OCT, US, FAG feautures
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Yooo!!!
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Why fiber rotator is needed in optical coherence tomography system? Currently I'm building a endoscopic OCT system from scratch and the fiber rotator will be used in the sample arm. Since I'm a newcomer in this area, could anybody offer an explanation for this device?
Thank you.
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Well, I never heard of OCT until this minute, but a fiber rotator is generally for selecting the axis of polarization entering (exiting) a polarization maintaining (PM) fiber. A couple minutes with Google and I see polarization sensitive OCT is a thing, so I think that is the idea. If the light source is polarized, the scattering tissue can change the polarization. Different tissues give different polarization effects, so being able to change what polarization you are observing provides extra tissue differentiation and contrast.
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Hi All,
I’m pretty new to cryosection so any advice or suggestions are greatly appreciate.
The brain tissues are fixed in 4% PFA and cryoprotected in 30% sucrose until they sink. Then samples were embedded in OCT and snap frozen in isopentane on dry ice.
The cutting temperature was set to be -18 - -20 deg C. What I noticed was that there are some irregular (not horizontal) wavy patterns fanning outward from the edge of the tissue (see picture), and when I tried to stretch them out, tissue will break apart. It is almost like the OCT expanded but the tissue didn’t (?) These waves on the OCT have caused lots of large air bubbles trapped underneath the tissue and wrinkles when I tried to pick it up using a slide. Any idea why this is happening and/or suggestions? Could it be the knife tilt angle?
thank you
stacie
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Hi Chen,
Your can try two things: First; reduce the volume of mounting medium. Since a difference in the density of the medium and tissue may also give such problem. For this you should either remount the tissue or cut out the extra medium carefully. Second, try increasing the temperature upto -15 to -18 but make sure difference in the temperature of the tissue and cryotome should be around 3 degree. Some cryotome models automatically take care of that i.e. there is only one temperature setting whereas in some you can set two independently.
These troubleshooting sequence you should follow is:
1. Cut out extra medium around the tissue.
2. increase temperature upto 15 (usually less then is not required and risky).
3. If your cryotome model supports make sure the temperature difference is around 3 to 4 degree.
4. Remount the tissue with less volume of medium and repeat these steps.
Hope it helps.
Best of luck
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I'm trying to stain mouse lung tissue IHC-IF
I fix in 4% PFA overnight immediately upon harvesting tissues
Then dehydrate/cryoprotect in sucrose gradients 15% to 30%
Then freeze in OCT media using liquid nitrogen
I'm sectioning on a cryostat at -20 degrees (5-10 um) and the tissue sections look to adhere well to the positively charged slides at this point.
Slides are kept at -20 until ready for IHC staining, but as soon as I try to remove the excess OCT with water or PBS, the tissue sections begin to wiggle loose. If I leave them in perm/blocking buffer for 30 minutes, they're half floating. And after leaving in primary antibody overnight, all tissues are floating.
How do I keep my tissues stuck to the slides? Is the PFA fixation before freezing affecting the charged slides from working like they should? Would a secondary fix step help- like a short 5 minute methanol fix?
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Dear Mayowa,
Baking the slices 1h @ 65 C works miracles.
Give it a try!
Cheers
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Hi,
I am trying to perform an IHC targetting a protein tagged with an Histidine tail. For this reason we ordered a recombinant Anti-6X His tag® antibody, which brings the following dilution recommedations depending on the application (image).
As you can see, the IHC dilution recommended is for paraffin-embedded samples and my samples are frozen and embedded in OCT( Optimal cutting temperature compound).
I will try to make different dilutions in parallel to optimize it, but, should I use dilutions near the IHC-P recommendation (1/16000) or to the one of ICC (1/100)?
*I thought It could be more similar to the immunocytochemistry because my samples have been directly frozen and the target protein should be more exposed than in tissues treated with paraffin, but I would like to listen to other opinions.
Thank you for the advice in advance.
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HIS tag produces a very strong signal. It is potentially further enhanced by unmasking when you boil paraffin slides in citrate or similar solution, even thought the HIS tag is either on C or n-terminus. With DAB detection, and 3-step antibody detection (primary -2ndary biotin -and avidin-HRP) there is substantial signal amplification, the dilution may actually be 1:5000 -1:10,000. When you do ICC, you do primary and then secondary-fluorophore, and no unmasking. Testing several dilutions of course is the best approach but i do recommend testing one at 1:5000 to 1:10,000 if you are using DAB as a substrate (more sensitive then fluorescence) and VectorLab or similar 3-step detection approach. Dont forget to quench endogenous peroxidase if you do HRP-DAB detection. Good luck
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There are so many different rodent OCT instruments and rodent fundus cameras on the market. Generally, the market is dominated by a Phoenix Lab Micron III/IVsystem however there little information about what performance you can expect from the different instruments. I was thinking of writing a review of the pros and cons of different OCT/Fundus machines so I'm looking for the OCT images of the mouse retina and cornea as well as fundus images to try to understand which instrument having the best resolution on the market. However, it is naturally quite difficult to have all the instruments tested. (I have data for Micron III, IV, OcuScience iVivo). Also, welcome your thoughts on OCT/Fundus user experience. It may result in the review paper, but too early to say. If you provide the OCT/Fundus images I'll contact you if we try to use it in the review.
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Dear, Brent A Bell Well, I have OCT images of the mouse retina on several instruments including Micron IV, Bioptigen, OcuScience with a few more it can be the foundation for the review. I hope to find few more collaborators to complete the set.
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Hi, I have OCT embedded tissue cut on Superfrost slides. However these slides are not Superfrost Plus, so apparently they are not coated. During IF all my sections are lost. How can I save them? Of course next samples will be cut on Superfrost Plus, but how can I save my current sections?
Anyone tried to coat slides with PLL with sections already on top? Or to fix them with PFA again?
Thanks in advance for any suggestion.
Cecilia
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Thenks for the advice.
Unfortunately the sections I am talking about are smaller than 1mm so it is really hard to move them. What I did instead was to define the region with sections with Pap pen, then incubate 30 min with PDL 0.1 mg/ml (sections were floating but remained in the region defined by the pen), wash with milliQ water and dry. Post fix with 4% PFA 15 min and rinse with PBS. Then I started my staining and, up to now, sections are still attached. Let's hope for the best!
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Hi,
I'm going to section vagus nerve tissue embedded in OCT, collecting 14 micron thick sections. What cryostat temperature is optimal for colleting these sections?
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-300C to -250C is optimum to use cryostat sectioning.
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I am analyzing the tissue damage to the mouse cornea tissue after incubated a kind of medium. The protocol is
1) mouse cornea was incubated in the medium for one day
2) embedding the cornea with OCT compound and snap frozen in liquid nitrogen
3) cryosection
4) HE staining
The control is naïve cornea that embeded with OCT, cryosection and HE staining. In fact, the thickness of incubated cornea should be thicker than the naïve one (3-4 times) from the optical coherence tomography. But the thickness of the incubated and naïve cornea from cryosection HE staining is similar. What the problem is? Should I use paraffin section? Why the thickness of incubated cornea is changed after cryosection and HE staining? Many thanks!
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use cryosectioning in which the tissue is immersed in cryoprotectant, frozen and then cryosectioned
fresh tissues were immersion-fixed by rapidly replacing the PBS with 10% formalin (Fisher Scientific, Pittsburgh, PA) for 1 minute at room temperature, taken out briefly for imaging and then returned to fixative for 24 hours. To study the effects of sectioning, the 24-hour fixed PPONHs were further processed, including immersing in a 30% sucrose solution for cryoprotection and embedding in Optimal Cutting Temperature compound (Fisher Scientific, Pittsburgh, PA) before freezing in liquid nitrogen (−196 °C). Frozen blocks of PPONH tissue were then cryosectioned coronally at 30 µm thickness (Leica CM3050 S, Leica Biosystems Inc., Buffalo Grove, IL)
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I am attempting to do mouse intestine longitudinal cryosections, but the sections after hematoxylin and eosin staining are not at their best (the villi keep breaking apart).
My protocol:
I rinse fecal matter out with PBS then fill with OCT (I used also a 1:1 diluted OCT with PBS).
I also tried to cut intestine lengthwise and sink in OCT - the villi curled outward, little better results.
I snap freeze in OCT and keep them in -80, cut at 10 um at -20C.
I tried to fixsome sections for 2 hours in 4% paraformaldehyde and then 30% sucrose overnight, but couldnt cut them - tissue too stiff after the fix?
Anyway, if you have any well-tested protocols for cryosectioning an intestine (not paraffin section!) or for the fixation of mouse intestine, please share!
Thanks!
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i am afraid you may not be using the best approach.
isolate tissue, rinse of course
fix with fresh 4% PFA (fresh PFA frozen at -20 and thawed ((less than 5-7 day old) is equally good
try fixing for 10 min, 30 min, 1h at room temp
wash 3 times 10-20 min each in PBs, using a shaker is better
cryoprtotect in 10% sucrose/PBS for 10 min
then add equal vol of 20% sucrose (to make it 20% weight/vol)
after 30 min to 1h remove 20%, and add 30% sucrose on PBS, keep until the tissue sinks to the bottom of whatever you use (flask, tube etc)
put tissue in cryomold (there are different shapes, if you need a long one, make it from a pierce of foil), add OCT, keep some 30% sucrose (about 20-25% v/v)
let tissue stay in OCT for 15 min or so
snap freeze in dry ice/100% ethanol bath, make sure you keep cryomold above ethanol surface or you will get a lot of bubbles made out of OCT (some do it in isopentane/dry ice and then add OCT, it does not work well for small tissue but works for brain)
keep at -80 until use
when cutting, optimize the thickness, for small fine pieces like villi you need 10-12 micron tissue. Optimize temperature inside cryostat, it impacts sectioning
if you plan to do H&E, postfix you frozen (well, lightly PFA-fixed) tissue with 0.1% glut/4%PFA, and maybe Bouin fixative, otherwise tissue falls apart (this is where one of your problems is)
i recommend avoiding H&E (it destroys tissue sometimes), and instead add a drop of CV (cresyl violet) for some 15-30sec, then wash with PBS, coverslip and image. This is more gentle stain for fragile tissue. However the bulk of fragility is caused, in your case, by no fixation at the beginning, and no postfixing before H&E.
if you are concerned about IHCs and impact of fixative on antibodies - it works, and you can do gentle antigen retrieval even on frozen tissue. PFA is a partially reversible fixative. Good luck
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Hi everybody,
I am working on cardiac fibrosis in mice and I am currently doing Masson's Trichrome staining. Since I still have problems with the staining, I hope someone has suggestions for improvement. The goal is to achieve a good contrast between the muscles and connective tissue (a strong red and a strong blue), but everything is currently purple. I have attached my protocol and previous pictures.
Thank you so much for your help!
Organ removal and fixation
• Open the chest and cut the right ventricle
• Insert the needle into the left ventricle and rinse with 1 ml 2 % PFA
• Remove the heart from the thorax and briefly rinse in 2 % PFA
• Press out the heart and weigh with a fine balance
• Rinse on the shaker with 2 % PFA for 1 h (The first thing we did was immunohistochemistry and an antibody only worked that way)
• Rinse on the shaker with PBS for 30 min
• Incubate overnight at +4 °C with 20 % sucrose
• Freeze organ with liquid nitrogen and store at -80 °C
Cryosections
• Cool the cryotome down to -30 °C (Leica CM3050 S)
• Store samples in the bottom of the cryotome
• Embed samples in Tissue-Tek® O.C.T.TM (Part of the heart that is cut is not embedded; OCT curls up when cutting)
• Cut slowly with 14 µm sections (the thickness also results from immunohistochemistry)
• Transfer the sample to a slide (Superfrost® Plus Thermo Scientific) using a fine brush
Protocol and Staining Kit: MASSON’s Trichrome with Anilin Blue (Article no.: 18156) from Morphisto, Frankfurt, Germany
• 04:00 Aqua dest.
• 10:00 Weigert's Iron Hematoxylin Solution (stock solutions mixed directly before staining)
• 05:00 Picric Acid Solution, alcoholic
• 00:05 Wash with aqua dest.
• 04:00 Acid Fuchsine - Ponceau (GOLDNER I)
• 00:05 Wash with aqua dest.
• 10:00 Phosphomolybdic acid 1%
• dry
• 05:00 Aniline blue (MASSON C)
• 00:05 wash with aqua dest.
• 02:00 ethanol 96% with 1% MEK
• 02:00 ethanol 96% with 1% MEK
• 01:00 isopropanol (2-propanol)
• 10:00 xylene
• 10:00 xylene
• 04:00 mounting medium
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usually, we fix over stain by using a differential solution, which is acidic alcohol 1% rinse the slides for 2-4seconds and makes your microscopic examination. check the concentrations of your stains, time of staining and you need to check the density of stating before you mounting the slides
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I need to devise a protocol for quantifying micro- and macro-metastases in murine lungs. We have several lungs stored in the deep freeze in OCT ready for cryosectioning. Is it possible to retroactively fixate these in Bouin's solution to provide contrast for surface metastases, then refreeze for cryosectioning and H&E stain to visualize micrometastases? Alternatively, is it possible to achieve the staining effects of Bouin's solution on frozen tissue either pre- or post-sectioning?
There are a lot of questions wrapped up in this, so please, any insight you have, do share! Also, if you have any references to provide for further reading, that would be much appreciated.
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Dear,
I have used frozen specimen for histology. But I was using 10% Neutral Buffered Formalin for the purpose. During my research, the availability of my research samples was at any time and more commonly in late night and it was very difficult to do dissection in odd hours (e.g. 2:30 am etc.). those specimens I was immediately storing in deep freezer at -20oC or below. Then I was thawing those specimen with lukewarm water to bring them at room temperature and in normal consistency. After thawing, I was fixing them in 10% NBF & Davidson's fixative for 24-36 Hrs. or maximum for 72 hrs. Then I was following routine histological processing and sectioning. I have used many staining techniques like H&E, Masson's trichrome, Verhoeff's elastic, Gridley's technique, Mercury Bromphenol blue for protein, PAS-AB for mucopolysaccharides, PTAH, Holmes etc. and did not face any problem with any stain I have used.
But I have never tried it with Bouin's fluid. So if you are planning for Bouin's fluid then you can try and I hope that there will not be any issue.
I am attaching file with some photomicrograph of my research to check the staining of tissue by above mentioned method.
With Regards
Dr. Mahendra
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I have OCT frozen kidney samples (not fixed) and I want to cryosection them to 5 micron sections. Kidney is a big tissue with large surface area so the minimum thickness I could reach is 15 micron. Thinner sectioning give sort of damaged sections or shattering although both the blade and glass slide are new and sharp. Is there any recommendation to be able to reach very thin sections using the conventional cryostat with no need for paraffin treatment of kidney samples?
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I routinely do 5 um cryosections of mouse kidney all the time at -20 temperature. Get it started with the anti-roll plate and then switch to a paintbrush to quickly section through the rest of it in one quick and smooth motion. It definitely takes a lot of practice.
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I am trying to extract intact nuclei from OCT-frozen tumor tissue samples. Does anybody have a protocol for doing so? I need as many intact nuclei as possible for performing ATAC-seq on them.
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I am trying to remove OCT from frozen tissue for DNA/RNA sequencing. My goal is to remove a portion of tissue from the OCT embedded tissue, without thawing or greatly disturbing the remaining tissue, and then using that portion of tissue for DNA/RNA sequencing.
Any help or suggestions are appreciated!
Thank you!
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If you leave OCT at room temperature, the OCT will unfreeze, however I do not know what will be the state of this DNA/RNA... However... You could try. Good luck!
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Hello,
I'm having some trouble with visualizing my IF in spleen tissue. I first found extremely high background and could not differentiate fluorescent cells and background. I then realized my stained tissue looks no different than tissue that was imaged prior to any staining or fixation. Any advice would be greatly appreciated!
More detail on tissue origin (not sure what's relevant...) 15um sections from rats perfused with 4% paraformaldehyde. Tissue removed and stored in sucrose after 24 hr in para. Tissue was frozen in OCT prior to cryosectioning and slide mounting. I've tried a few recommendations with no success (glycine, ammonium chloride and trying overnight fixation).
Thanks in advance!
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Brady! Is the auto-fluorescence laser/filter cube-specific? If so, you may want to switch secondary antibodies and avoid extensive trial-and-error problem solving.
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Looking for public database/source of en face OCT angiography (OCTA). Anybody know where can I find OCTA en face images? I have several b-scan sources but cannot find en face OCTA images. Thanks
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Hi, I hope this answer is not too late. You mat find datasets at:
and here:
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I am trying to sectioning fresh frozen mouse testis, but the H&E doesnt look good. I am enbedding in OCT and cutting in cryostat at 10 um. The temperature of the blade is -15C and the sample is -10. After that, I tryied to fix in bouins solution or formalin for 1h. But the morphology of the tissue doesnt look good.
Any tip?
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We are processing pig lungs for ICC. We inflate the lungs with air, then cut a piece and fix it in paraformaldehyde. The next day, we change it to 30% sucrose and wait until the tissue sinks to embed it in OCT. However, most of the lung pieces do not sink after 3-5 days. Does someone have had the same problem with lung tissue? Should we be using a different sucrose concentration or a different cryoprotectant?
Thank you so much!
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I am trying to perform HE staining of small intestine. I used OCT solution to make sections.
After HE staining when I look my slides under microscope, I do not get intact villi. In much of the cases, villi are broken. Does anyone can help me?
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IMHO, the problem is the incorrect orientation of the block when cutting. You have slice the block perpendicular to the axis of the digestive tube fragment. Then the chances of getting whole villi in the slice along their entire length significantly increase.
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Due to COVID-19, our lab is shutting down. However, we have rats that have been transfected with AAV5-CaMKII-ChR2-mCherry virus and fiber optic implants. To make life easier for the vet techs, we may sacrifice the rats.
Normally, I would slice and stain them within a week of perfusion, but I want to minimize the time I go into the lab.
What is the best way to preserve the brains for about 2 months?
They will be perfused with 4% PFA, then put in sucrose PB. Normally, I would slice with cryostat, stain with primary anitbodies anti-RFP and anti-NeuN, then secondary antibodies conjugated with AlexaFluor. My proteins of interest seem pretty robust, so could I just leave them in sucrose solution at 4C? My biggest concern is ice crystals tearing the tissue if I freeze and leave it for months, although I haven't had issues when I've stored brains in OCT in -20C freezer for 1-2 weeks.
We have -80C freezer and OCT, but NO liquid nitrogen or isopentane.
Any advice is greatly appreciated.
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Residual PFA is supposed to be one of the main things that affects the flurophores, so rinsing the brains with PBS several times could help before putting them in sucrose.
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The retinal scan area of an OCT is derived by calculating the linear distance on the surface of the retina subtended by a fixed field of view using a defined axial length. This is 23.82 mm for the RTVue XR Avanti system (Sampson et al. 2017 - https://doi.org/10.1167/iovs.17-21551) and 24.46 mm for the Cirrus HD-OCT 5000 (Shpak & Korobkova 2020 - https://doi.org/10.1007/s00417-019-04513-w).
Does anyone know the defined axial length for the Topcon DRI OCT-1 Triton?
Many thanks in advance,
Zoran
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The default Axial lenght of Topcon DRI OCT Triton is 24.39mm
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I have spleen samples from infected mice which are currently frozen in OCT. Ultaimtely I would like to use immunohistochemistry to image spleen slices and see the ROS within different infected/uninfected macrophage subtypes. I have found a protcol that allows fixation of cells whilst maintaining the ability of DCF to stain ROS (using 10% Methanol), so the IHC itself shouldn't be an issue. My question is: are ROS likely to be stable inside macropahges within spleens frozen in OCT, or will they break down?
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Dear Ryan,
I completely agree with Antons comment.
ROS measurements are done routinely in our lab mainly with different ex vivo cells from the innate immune system, but it took a long time to set it right.
One of my concerns about ROS level determinations either with Microscopy or FACS is, that they are only snapshots of a rapid response. Macrophages, neutrophils and PBMCs react very quick (at least after infection and PMA) with ROS production (peak after 30 min) and end approximately after one and a half our. I would always recommend ROS measurements in a plate reader with kinetics instead of making "ROS-Snapshots". Moreover you do not encounter the problem of fixating the ROS probes.
There are ROS probes, wich I would not recommend.
The problem with luminol is, that it freely diffuses over cell membranes. So by using this substance you get a "overall ROS measurement". The same goes with the generally used DCF substance H2DCFDA, which a lot of people use. This also freely diffuses over membranes, so with that DCF probe you only detect "overall ROS" in the cell.
For both ROS probes derivates exist that are not cell permeable (isoluminol) or are retained only in the cytosol (6-Carboxy-DCF). The protocol for Isoluminol also works fine with Amplex Red (measures specifically extracellular H2O2).
Not all cells produce ROS in the same amount, in all compartments with every stimulus. What induces ROS in one cell type might not induce or even decrease ROS production in another or perhaps with other kinetics. If you know a stimulus that works in your other experimental read-outs and this stimulus does not induce ROS production with the same concentration, then there is simply no ROS production that can be detected.
For any questions, informations or technical details for ROS measurements and how these measurements look in a plate reader (in macrophages with different stimuli, scavengers and in different compartments), I would like to highly recommend our paper. 70 % of it is more or less about ROS measurements in a plate reader. The protocols described there worked in our hands with macrophages, neutrophils, human PBMCs, Microglia and MEFs.
Mitochondrial reactive oxygen species enable proinflammatory signaling through disulfide linkage of NEMO. Science Signaling (2019)
For a overview of ROS in general I can recommend:
Reactive oxygen species (ROS) homeostasis and redox regulation in cellular signaling (2012), Paul D.Ray, Bo-WenHuang, YoshiakiTsuji
ReviewROS Function in Redox Signaling and Oxidative Stress (2014), MichaelSchieber, Navdeep S.Chandel
Please feel free to ask any further questions or add your experiences.
All the best,
Marc
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Hi
I have neural tissue that I am embedding in OCT freezing medium and cutting to 18 microns on a cryostat. I store my slides at -80 until staining. What I noticed during my last stain is some pieces of the tissue fall off the slide and float in the coplin jar. I have also seen pieces fall off when I drip antibodies on the slide. Why does it happen and how do I prevent it?
Thank you!
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You know how you have your water bath, right? So, what you do is you sprinkle powdered gelatin on the heated water - enough to cover the surface, but not fully cover it so. It will dissolve. Then, you dip you slides with the specimen already on it, covering them in the water, and then you put them on a slide warmed to dry up. If you don't trust you slides in the first place, then you use that water instead of your usual water bath when picking up sections (which is how I was shown to prepare water baths in the first place). I know, it's so silly but it works!
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Hello. I want to do some image analysis on OCT blood vessl images. Given the contour of the adventitia, how can I calculate the center of the main blood vessel?
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You can use image-j (N.I.H) software.
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I have flash frozen some brain slices in OCT with isopentane, cooled by liquid nitrogen
I have mounted some on slides using a cyrostat however I am finding that the tissue dissociates from the slides very rapidly.
I just wondered if anyone had any advice? It was deemed unsuitable to fix the tissue with PFA before storing in the -80
Thanks
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I agree with Martin Schol , I use Superfrost Plus™ Slides by Thermo Scientific which electro-statically attract frozen tissue sections. I mount with them directly from sectioning in the cryostat and the tissue
is adhered to it properly.
Good Luck!
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Hi all - how long do you trust your frozen slides in the -80 C? I cut half a block at 6 um, but the staining core closed for break before I could submit them for immunostaining. We fix whole tissues in PFA, then freeze in OCT, then section.
Thanks!
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I would venture to say that as long as they are stored properly in a well-maintained freezer with manual defrost that they should be good indefinitely.
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Hi,
I want to freeze D0 pups n OCT blocks while they have been cut through the median plane to get the symmetrical images of different organs in each half of the animal body.
I just not sure if I can get a sagittal section of such a big tissue block using cryostat?
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Thanks for your comments. I felt that it is not the best idea. My goal is to get the dorsal ganglia in D0 pups and it's very hard to dissect them. Also to get a good orientation.
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I am trying to do Laser Capture Micro-dissection on samples that have been frozen at -80C, for many years. My purpose is to isolate specific cell type and isolate RNA for RNA-seq experiment. I have read some reports saying that it is not possible to use OCT if your tissue is already frozen. My question now if this is just a technical issue that can be solved or some serious problem with no solution?
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HELP!! my tissue sections have thawed overnight in OCT medium, can I salvage them? HOW?
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Hello all,
I have an issue for cryosection during these days. My samples are all mice brain tissue. The size of these tissues is all around 1~2 square millimeters. I found each section has the same cracks at the same place. Could anyone can help me to figure out what step is wrong for the cryosection?
Step1. I fix the brain tissue sample by 4% paraformaldehyde at 4ºC overnight.
Step2. Add 15% sucrose shaking 10 min, change to 30% sucrose stay at RT temperature overnight.
Step3. Put the sample in OCT around 2 hr.
Step4. Frozen the sample in OCT at -20ºC for 2 hr.
I attached a picture for the cryosection, you can find many cracks on the section. How to avoid this problem?
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Iagree step 1 is fine. However, in my old lab we put brains in 15% sucrose at 4ºC overnight, then next day into 30% sucrose at 4ºC overnight, no shaking for either. Then we would put brains in OCT in cyrovials and immediately put onto dry ice to freeze, then transfer to -80ºC for long term storage or -20ºC for short term storage (if cryosectioning was happening soon, or you can cryosection immediately once OCT is frozen).
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I would like to de-embed fixed tissue that are stored at -80° in OCT resin.
Final aim is to perform a whole mount immunostain with.
Does anyone have experienced such a procedure?
I would appreciate feedback or troubleshooting advices.
many thanks
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OCT is water soluble. You shouldn't have any issues washing it off and subsequently staining following a normal permeabilization/blocking procedure. The whole mount staining procedure is, however, tissue dependent (wash and stain times/concentration of TritonX or other detergent etc.). More details are needed to inform on the staining side of things. Good luck!
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My question pertains to human heart tissue previously embedded and frozen in OCT.
We have recently cut sections of varying thickness (100-1000 um) for processing and staining while free-floating. The sections were cut on a cryostat and transferred immediately to formalin for fixation. However, while the thicker sections remained intact, the thinnest section, ~100 um thick, seemingly completely disappeared following 2 hours of fixation. Has anyone previously observed the dissolution of very thin tissues in formalin?
Thanks.
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To me even a 100um section is "thick". You should be able to do free floating sections 20-30um with no problem. I assume your fixative is 10% formalin (aka 4%formaldehyde) either 10% NBF or 10% formal saline. If they are falling apart is it due to slow freezing and ice crystal artefact? Your tissue may not be "fresh" as well and I assume may have some "pathology" associated with it?
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I am trying to analyse some OCT pullbacks using Osirix, having never used the software before. My aim is to measure CSA at various points of the pullback, and also to semi-quantitatively analyse the plaque characteristics. Any advice will be very hepful
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I think OCT image in the cardiovascular field has been defined by DICOM format.
You should modify open-source version of OsiriX in order to read the format.
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Can I store my samples (fixed in 10% neutral formalin) in OCT at -81
˚C. How many times can I frost/defrost there samples withour their damage?
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Yes, you can store the fixed sample in OCT in -80C freezer.
One time is always preferred. The more times the sample is frost/defrost, the more damage it will cause. Depending on the sample type, skin sample is more robust than brain sample. The brain sample will be fully damaged after 2 frost/defrost cycles.
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Hello experienced people in IHC/IF,
I am doing an IF experiment to stain free-floating, 30 μm, mice hippocampal brain slices for a membranal protein. However, when I made sections using a cryostat, I found that my slices rolled up upon transfer to PBS. I used 1% PFA+1% sucroce in 0.1M PB for light perfusion of the brain (as per the protocol for my protein of interest). I embedded the brain in OCT before cutting and used the anti-roll glass while sectioning. The slices looked fine until I put them in PBS. I was wondering if anyone knows how to unroll them using any known alternative methods.
Thanks for your help/suggestions in advance.
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Hello Rinki Di,
Thanks a lot for such a detailed explanation. I believe these tips will definitely help me in processing the brains in the near future.
And you are absolutely right about using the vibratome. I had very good results with vibratome in regard to sectioning in the past. Unfortunately, as for now I have already used cryostat to process the brains.
Thank you for your time and assistance.
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I am having difficulty getting specific binding with my rat spinal cord immunofluorescence. My protocol is as follows:
1. Remove spinal cord and quick freeze in TissueTek OCT.
2. Cut tissue on cryostat, laying slices directly on gelatin coated slides.
3. Acetone (-20 C) fixation for 10-20 minutes.
4. PBS wash twice, 10 minutes each.
5. Blocking for an hour in blocking buffer (5% FBS, 0.2% Triton X-100, PBS) at RT.
6. Drain blocking buffer and incubate primary antibody mixed in blocking buffer for 2 hours at RT (have also tried overnight at 4 C).
7. Rinse slides in PBS 3 times, 10 minutes each.
8. Incubate secondary antibody and DAPI (0.5 micrograms/mL) in blocking buffer for 1 hour at RT.
9. Rinse slides in PBS 3 times, 10 minutes each.
10. Dehydrate with 50%, 80%, 90%, and 95% alcohol.
11. Mount with Vectashield mounting medium.
I have tried this method with many different primary antibodies, all validated for IF. The secondaries are from Vector Laboratories (Fluorescein and Texas-Red conjugated mouse/rabbit anti-IgG). My DAPI staining is not uniform across my slices; there are areas that you can see the nuclei very well, but others on the same slice are very dark). Any help would be appreciated!
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Perfusing in PFA and immersing in sucrose was the method my PI previously used, but this new method was suggested and we are trying to get some results since it is easier. But I may just suggest to my PI that we go back to the original method. Thank you both for your answers!
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I want to use TUNEL (In Situ Cell Death Detection Kit (Roche)) to detect apoptosis in mouse lung tissue that has been frozen and sectioned in OCT. Suggestions for how to prepare tissue prior to using the kit?
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Hi,
I have snap-frozen (isopentan-bath) mouse kidneys stored at -80°C and embedded in OCT. How can I de-embedd them to use for a Nephrin ELISA?
Thanks.
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I am using laser diode centered at 840 nm as the source. The experimental setup is similar to the figure shown below. The beam splitter cube splits the beam into reference and sample. The coating for all lenses and the beam splitter is for the specified wavelength. The objectives in both the arms are not coated for 840 nm wavelength but has transmission of 60 % at that wavelength.
When I image the sample on the camera, the reflection from the back surface of the beam splitter is prominent. Rotating the beam splitter deflects the beam on the objective but does not necessarily remove the back-reflection (ghost image) from the beam splitter.
How to get rid of back reflection?
Which beam splitters are generally used for Full-Field OCT or Linnik based interferometry systems? I have gone through few papers and they use the similar beam splitter as I use- Non-polarising cube beamsplitters and they don't face the ghosting problem.
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If the the optical path length's between sample arm and camera is not a problem, then you can try introducing a wedge (optics) in the camera leg and relocate the camera to reflected spot from the wedge. Because of wedge angle the back reflected spot can be separated considerably from the main spot. By adjusting the distance between camera and wedge can also help you in separating the beam considerably. Disadvantage of the scheme could be low intensity of spot, but i believe that can be managed by gain features available in camera or with associated electronics for the camera.
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Hypertensive patient presented with feature of macular BRVO without CME which was further confirmed with OCT. What is the ophthalmic management in this case. Systemic ix and referral to internist was given
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As we don't have mfERG we will do 10-2 at baseline and will monitor with amsler grid. Thanks alot to you all for this fruitful and debatable discussion
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I have been working on cryosectioning adult zebrafish with large tumors in their abdominal region, but I am getting a significant degree of holes/tearing in the sections as they come off the block. I'm not able to fix the fish before sectioning as the experiment the sections will be used for requires fresh-frozen tissue.
My method: I am euthanizing the fish in ice water and then immediately flash freezing in OCT/dry ice/ethanol. I am leaving the blocks at -80 for at least 24 hours before sectioning, which seems to help with the tearing issue somewhat.
There is not much information on cryosectioning zebrafish online (that I could find) so I was wondering if anyone has any tips?
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Hi,
Please, have a look at these two documents:
Also, did you coat the mold with an OCT layer sitting in some dry ice prior to covering the zebrafish with OCT completely and keeping it on dry ice?
You might have some ice crystals forming if you place the freshly eutanized fish directly in the mold with OCT on dry ice, since freezing is a bit slower.
If the above issues have been avoided, have you considered mucin as another possible culprit:
as mucin expression pattern and composition is altered in cancer animals.
I hope this helps :)