Science method

Microscopy - Science method

The use of instrumentation and techniques for visualizing material and details that cannot be seen by the unaided eye. It is usually done by enlarging images, transmitted by light or electron beams, with optical or magnetic lenses that magnify the entire image field. With scanning microscopy, images are generated by collecting output from the specimen in a point-by-point fashion, on a magnified scale, as it is scanned by a narrow beam of light or electrons, a laser, a conductive probe, or a topographical probe.
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Fellow Researchers, I need some wisdom.
I am currently performing an experiment involving CaCo2 cells seeded on a Transwell. I don't have experience with such assays, therefore please forgive me, if my questions are somewhat ignorant. I've searched ResearchGate and found some answers in this discussion:
(23) Can transwell-cultured Caco-2 monolayers be imaged with BD Pathway confocal microscopy while keeping them in transwell? (researchgate.net)
However, I wonder - is there a possibility to observe the Transwells while on a plastic plate, without the necessity of transferring the Transwells to a glass dish? Additionally, I use the type of Transwells that stand on the bottom of the well, not hang on the borders of the well, therefore I am not sure if this solution would work in my case. My idea was- if I used a non-inverted microscope, could I maybe see my cells in the Transwell? I understand that after seeding, my cells need 21 days to differentiate and form a monolayer, however, how can I check that they are in fact attached to the Transwell and growing if I have no way of seeing them? Is staining with some viability dye the only way? I am afraid of a scenario in which I wait 21 days for my cells to differentiate while in fact – they didn’t even attach?
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Hi Ariadna,
I work with primary lung epithelial cells in transwell cultures, but I can give you some ideas on how I do my cultures.
During the differentiation and maturation phase I image my transwells on an inverted brightfield microscope. whether this will work on your microscope depends on the working distance of your objectives. Our confocal certainly doesn't have the working distance needed for in-situ imaging of the transwells.
For my confocal imaging, I fix the membrane and cut it out, imaging it as a whole mount on ordinary glass slides. Were you wanting to do IF images at different time points?
Also worth noting (at least for my primary lung epithelia) that the cells should grow to full confluence in a few days, so make sure you seed sufficient cells (I seed 1.5e5 cells per 12mm insert). It's the differentiation that takes time.
If you're just wanting to check that there are cells present, they should be visible on an inverted brightfield microscope. I of course have the advantage that I can see beating cilia and mucous production under the brightfield as signs of differentiation.
Another measure of cell-layer intergrity and barrier function could be trans-epitheial electrical resistance (TEER). This can be measured over time, without killing your cells. There's a neat paper on constructing your own device to measure TEER if you don't have the budget to buy a commercial device:
Hope this helps!
Sam
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Hello,
I'm trying to develop a solution that can do particle analysis (count, diameter, area) on dark-field optical micrographs of semiconductor thin films (samples attached). I have been using ImageJ so far. But I'm facing some problems: my code needs to process hundreds of images, and ImageJ takes a long time to run. Moreover, the results heavily depend on the choice of algorithm and various other parameters.
I am therefore looking to develop a solution in Python. I'm now considering deep learning using PyTorch. I am wondering if there are more straightforward solutions out there. Moreover, are there any publicly available databases available for training the neural networks, or is creating them myself the only way?
Thanks,
Pouyan
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There are dozens of fully functional programs for solving such problems (I will not enumerate, this will be advertising). I met the first of them in the mid-1990s... Is there a need to develop a new program, especially since for interpreted Python, the low speed and higher memory consumption of programs written in it are well known compared to similar code written in compiled languages, like C or C++? But it is popular and fashionable, that's a fact... for professional programming, because it allows you to produce more code (especially in large companies and for mass applications).
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The use of resin in microscopy has been expanded since the discovery of its use in 1950s and 60s. What are its other uses now?
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Resin/plastic is used for electron microscopic studies and to examine hard tissue, such as bones.
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Dear colleagues,
Could we use scanning electron microscopy for investigation of the surface morphology of piezoelectric ceramics? May there arise some problems with piezoelectric effect during microscopy? Could we prevent such problems if we use conductive coatings and supports?
I would be grateful for your assistance
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Another work around, if you must take images and you do not want to use coatings as it can mask features you want to see, of a non-conductive sample is to place a (thin) strip of conductive tape, usually carbon tape or copper, across the sample surface. Ensure that at least one end of the tape is affixed to the stub (grounded).
Often, you can image near the conductive strip as it conducts the electrons away enough so you can get very good images if you are adept at SEM. I have used this quite a lot on ceramics, it should also work on other materials.
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Hello everyone!
Looking for some useful tips for fluorscence/confocal microscopy of plant root tissue.
I am mostly working with the root tissue of Arabidopsis thaliana where I take confocal/fluorscence images of plant root samples expressing proteins of interest with GFP/mcherry tags also often with Propidium Iodide/DAPI stains. Usually, I don't fix the tissue and take images as such and here are a few problems I face most of the time and I will be glad if you can advise me some suggestions to fix or minimize the following issues-
1.The amount of protein expression in various layers of root tissue is variable ranging from low to very high expression so in a Z-stack often I get highly staurated layers or if I adjust laser power/analog gain to minimize that oversaturation I end up losing signal where protein is less expressing. What should I do in such case?
2. how do you prepare live tissue for imaging, like if you use any specific buffer that does not change the physiological state of the plant tissue but helps in maintianing the GFP fluorescence for a longer duration?
3. Also, no matter how carefully I prepare the slide I never get the root tissue lying uniformly over the slide which means I get regions with good focus and in perfect plane wherease at some regions it gets blurry, out of the focus, how to avoid it?
4. I too have problem with DAPI staining of the root tissue as in most cases barely I see any nuclei but most often I end up with images showing DAPI at the cell borders just like Propidium Iodide.
Thanks a lot in advance for all the incoming beautiful suggestions.
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I work mainly with mammalian cells and have no clue about the plant cells but I feel this could help you with your DAPI problem. DAPI is not great at staining a live nucleus. DAPI most often cannot enter cell membrane/wall and staining requires fixation and permeabilization. I suggest you try Hoechst 33342 which has similar excitation and emission wavelength so you don't have to change your protocol.
Please note that this works for mammalian cells and may or may not work for plant cells but something to think about.
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Dear community,
I am facing a problem with Fe(III) minerals that I cannot dissolve with my usual approach using oxalic acid solution and iron EDAS. I am studying bacteria that oxidize iron. In order to do some analysis, I need to remove the Fe(III) minerals formed in my system. In my current setup with ziro valent iron as source, the formed minerals do not dissolve easily. Adding, for example, 6 M HCl would probably also destroy my cells, so I would not be able to examine them. Therefore, I am looking for ideas to solve this problem. This could be a special fixation method, or reducing agents that dissolve iron without being extremely acidic.
Thank you very much for your comments!
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See the review of Voelz et al. 2019 (10.1021/acsearthspacechem.9b00012) for different iron mineral dissolution methods.
It depends on which iron minerals you want to dissolve but a dithionite exraction seems promising to me as long it its not magnetite .
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During AFM imaging, the tip does the raster scanning in xy-axes and deflects in z-axis due to the topographical changes on the surface being imaged. The height adjustments made by the piezo at every point on the surface during the scanning is recorded to reconstruct a 3D topographical image. How does the laser beam remain on the tip while the tip moves all over the surface? Isn't the optics static inside the scanner that is responsible for directing the laser beam onto the cantilever or does it move in sync with the tip? How is it that only the z-signal is affected due to the topography but the xy-signal of the QPD not affected by the movement of the tip?
or in other words, why is the QPD signal affected only due to the bending and twisting of the cantilever and not due to its translation?
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I agree with Annemarie Honegger.
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Hi all!
Please could someone tell me what is this structure that stains PGP9.5- positive on a hair in rat skin? (see picture)
I am new to skin morphology and textures, and even though I am starting to understand the pattern of innervation, I found a structure that is unfamiliar with me.
It looks like cells with some kind of extended structures / axons? But I do not know of any neuronal cell type that surrounds the hair? It looks like a pericyte but it can't be.
The picture is of Rat dorsal skin, blue = dapi/autofluorescence, green = pgp9.5 (or cell-specific autofluorescence ?)
Does anyone have an idea? Schwann cells?
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I'm not a skin expert, but my first guess (based on the morphology) would be Langerhans cells. And after a quick Pubmed search they also seemed to be also PGP9.5 positive.
Kind regards
Soenke
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Hello everyone,
I would like to do some light microscopy to observe cells that have grown on an insert. Do you know if it is possible to fix the cells and stain them with ORO directly on the insert, then arrange the insert between slide and coverslip? And if it is possible, how would you do that?
Thank you very much to anyone willing to help =)
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Yes, baring some unknown condition, your idea should be doable. I would suggest that cutting out a segment of the membrane containing the cells would be a good way to start. Place the membrane in a Petri dish, cell side up. Stain the membrane drop wise, being careful not to wash off the cells. Forceps can be used to carefully allow the stain to run off. Do a single rinse with water, drop wise. Then observer the stained cells.
This may require a little practice to get it right, but should be doable. I have done similar staining techniques. With practice, this should work.
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Some of the diatom species i cultured undergo frequent auxosporulation, but i have not been able to find sperm in the LM (1000x / 1,4). It might be an asexual process, but maybe sperm is infrequent or inconspicuous… Advice for SEM is also welcome.
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In the light microscope I have, seeing spermatogonia is not a problem when looking at salmon testis. Seeing individual mature sperm cells is not possible. You see clumps of mature sperm cells but there's no way of seeing individuals unless you go to the TEM or SEM. If the sperm in diatoms is infrequent, they will be inconspicuous by light microscopy.
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I am trying to do some fluorescent microscopy on E.coli and P.aeruginosa cells after 24h treatments with compounds. I am using propidium iodide (molecular probes) and SYTO 9 (Thermo) in the following way:
1. Add equal volumes of each dye to 100ul of 20% glycerol
2. Add 2ul of the dye to mixture to samples in microtiter plate
3. Cover with foil and incubate in dark for 15 minutes
4. Pipette 10ul samples onto slide and view under fluorescent microscope
When I come to view my cells under the microscope, I can only see them under light microscopy and when I switch to using the fluorescent filters, I see the same cells in both filters and none of them are fluorescing green or red.
I tried just adding each stain separately and the fluorescing cells in each filter can be seen but not when I add it is a mixture of both dyes.
Could someone assist me?
Michael
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Dear Michael,
Kindly optimize your mounting media. Because fluorescent dyes are susceptible to many of the components, the fluorescence ability of PI and Syto9 is quenched by mounting media or due to some non-specific reactivity.
Also, I suggest you kindly do not use glycerol for the dye to dissolve your dyes as mentioned by the vendor (Thermo Scientific). Glycerol may cause a quenching effect on your dyes.
Go to this link you will find the Live Dead Bac kit and protocol try your experiment accordingly. Hope you will definitely find it helpful.
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Dear environmentalist in Bangladesh,
I would be happy to know where I can get the FTiR microscopy facility and the already developed protocol for micro-plastics characterisation in biological samples in Bangladesh?
Also suggest any transcriptomics marker to analyse in fish and molluscs.
Thanks in advance.
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This article might help you to MP analysis. You can get a detailed idea from the corresponding author of this paper.
Jabed Hasan, can you please help him to characterize the MP.
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Hi, is anyone here using a light-sheet microscopy? What commercial brand would you advice?
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Zeiss products good in microscopy, I have used one stereo
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We are planning to study inflammatory cells in mouse bronchoalveolar lavage fluid. Briefly, we would perform the bronchoalveolar lavage, collect the fluid, centrifuge it, freeze the supernatant at -80 degrees C with proteinase inhibitors. Then we would lyse erythrocytes with ACK lysis buffer for 2 min., dilute the ACK buffer with PBS, centrifuge again.
I have two questions:
1. Should we add EDTA to prevent cell clumping?
2. How should we perform the Wright-Giemza stain? We do not have a cytocentrifuge. Can we just resuspend the cell pellet in a small ammount of PBS and add the cell suspension onto a slide to dry out and stain? Maybe someone has a similar protocol that you could share?
Thank you!
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You need to have some method of centrifugation for the cells to stick to a slide and get a bit flattened out so that their features can be evaluated microscopically. If you don't have a cytospin, there is another technique which involves a basic centrifuge. Use cylindrical flat-bottom tubes, inside which you place a round slide coverslip of just under the diameter of the tube. Then put the BAL fluid in the tubes and centrifuge. The cells will stick to the coverslip, which you then remove from the tube and stain with your method of choice, and then mount the coverslip cell-side down on a standard microscope slide.
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I work on plants of the genus Crassula, which have succulent leaves with usually thick cuticle. For microscopy, I'm currently sectioning resin-embedded samples in an ultramicrotome. In a nutshell, the protocol I'm using is as follows:
1. Fixation in formaldehyde (4%)
2. Ethanol dilution series to dehydrate
3. Embedding in LR white resin (medium grade) + resin polymerisation
4. Sectioning in an ultramicrotome using glass blade
I'm obtaining sections that are 1 or 2 μm thick. The problem is that, even though most of the sample looks fine, while sectioning the resin tends to detach from the cuticle and rips off many epidermal structures. This leads to either broken epidermis or greatly distorted, and I also lose most of the details of the indumentum.
Has anyone had a similar problem while sectioning resin-embedded samples with thick cuticles? If so, what would you recommend that I change to prevent this from happening?
Thanks :)
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Alistair Leverett Var St. Jeor Sabine Dürr Thanks so much for your suggestions! I actually cut the succulent leaves into 2-3mm sections before the dehydration and embedding in resin, to make sure that the resin will penetrate the tissues. So most of the surface is already exposed mesophyll, so that reasin can penetrate. When looking at the sections, the appearance inside the tissues seems homogenous, which leads me to believe that the resin penetrates properly. It is only the cuticle itself which sometimes detaches from the resin and tears the epidermis in the process. I will reassess the protocol that I'm using, maybe I can make some adjustments following your suggestions. Thanks again :)
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I have these oxide/metal/oxide sandwich samples that the thickness of it reaches below 100 nm in the areas indicated in the picture. Those are oxide/oxide regions. I am just concerned about structural studies in the aforementioned areas. The thickness of normal areas of the sandwich is around 5 micrometers. The question is:
1. Can I take it directly in the TEM sampler and just search for those areas without any sample preparation?
If not, what kind of sample preparation do you suggest(not expensive methods like Ion milling and FIB?
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I am happy to announce that the TEM investigation of the sandwich samples was successfully done without any pre-requisite procedures. The results for Al-Mg and Al-Mg-Be sandwich samples is now presented in the following paper:
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Solvents for the immersion oil are carbon tetrachloride, ethyl ether, Freon TF, Heptane, Methylene Chloride, Naptha, Turpentine, Xylene, and toluene.
What is the best of these to clean the surface? Toluene? Heptane? I'd like to stay away from more dangerous chemicals if possible and have something that evaporates easily.
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Methyl alcohol is the best
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Recently my lab has run into an issue when using Alexa Fluor 647 on mouse brain tissue (PFA fixed/frozen IHC). When viewed under an epifluorescence microscope (Y5 filter cube: Excitation: 590-650 & Emission: 662-738), the telltale staining pattern for microglia can only be seen AFTER viewing of the DAPI channel and quickly loses intensity until the signal fades into the background. Exposure/viewing with the Y5 filter cube without first exposing to UV yields very poor signal; so poor, in fact, you would believe the staining was a complete failure. It's as if the 647 is being excited by UV instead of the intended ~650nm wavelength, but properly emits the correct wavelength. We use Vectashield Antifade when mounting and the signal always reappears perfectly after UV exposure, so it doesn't appear to be photobleaching. When setting up a confocal microscope to view 647, there is no discernable signal (will test at UV settings soon).
Has anyone seen anything similar with their own IHC staining? Can anyone explain this phenomenon? Would switching to Dylight 650 potentially be a better alternative, or do red leaning fluorophores have issues? I appreciate any and all input and thank you in advance!
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Dear Amanda,
indeed we also had to figure out that Vectashield is not compatible with Alexa647 staining. We use DAKO fluorescent mounting medium or FluoromountG with DAPI.
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I want to analyse cross-section samples by SEM microscopy whose constituent layers have been applied by dip-coating using SS and glass as substrates. However, I do not know if I would have to prepare the sample in a special way for its analysis and if it is the case, how should I previously prepare the sample without altering the applied layers? I have already tried to cut the glass samples using diamond tip cutter but the cut was neither clean nor precise.
Thank you for your help.
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If ion mill (as described by Pierre Caulet ) is out of reach, you can use metallographic polishing. To preserve thin layers it's better to make specimen "sandwich": with thin epoxy glue two pieces of specimen with substrate on outside. Embed, polish.
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Hey,
I am working on nematodes and my interest is in strongyloides stercoralis. I have obtained larvae from fecal samples of goats and sheep by kato katz culturing method aka charcoal culture method and extracted larvae on 3rd-20th day. Now I am facing difficulty in differentiation of strongyloides stercoralis larvae and other hook worms larvae. I am trying to confirm them by microscopy and focusing on tail morphology of larvae i.e bifurcation in the tail of the larvae but it’s quite difficult and time taking process as for tail ive to see all the larvae at 100X (oil immersion). I am sharing some pictures (10X, 40X and 100X) of the larvae in hope to find some suggestions and materials for morphological identification.
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Dear Nadir,
This paper can be useful for you.
Please consider the possibility you are dealing with other species from goats, such as Strongyloides papillosus.
Hudson
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Hello, I am a student currently working on an MLA for a multi-focused camera. One question arises and I ask a question.
Multi-focused plenoptic cameras from most of the papers adjust the focal length by setting the three lenses the same diameter and different RoC. However, my MLA is made with different diameters of lenses and different RoCs.
Does this cause any imaging problems? Thank you for your help.
I attached the fabricated MLA's Optical microscopy picture
-sincerely-
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I created a quantitative image of the image you want for you with an algorithm.
I hope it helps you solve your problem.
with the best wishes
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A hologram is made by superimposing a second wavefront (normally called the reference beam) on the wavefront of interest, thereby generating an interference pattern that is recorded on a physical medium. When only the second wavefront illuminates the interference pattern, it is diffracted to recreate the original wavefront. Holograms can also be computer-generated by modeling the two wavefronts and adding them together digitally. The resulting digital image is then printed onto a suitable mask or film and illuminated by a suitable source to reconstruct the wavefront of interest.
Why can't the two-intensity combination algorithm be adapted like MATLAB software in Octave software and create an interference scheme?
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I would like to perform reflected light microscopy to image the surface of a sample only, but only have access to a Zeiss Axiovert inverted light microscope. Is there any way to modify my scope to do this?
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Observing a covered slide in reflected light is certainly not a standard procedure, but experiment. In a bright field at lower magnification, only a reflection from the cover glass is essentially visible. However, it is possible to experiment and combine techniques in various ways, sometimes even slightly incorrectly. However, most procedures require good quality, preferably with a cooled camera, because the lights are low. For example, at higher magnifications (40x or 100x objective) with immersion, something (surface structure of spores) can be seen in polarized light. As Guenter Grundmann mentions, dark field or phase contrast works better.
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For training a network I'm searching for a dataset of complex bio-images. I have seen some datasets for cell-classification, but these are usually simply amplitude-only samples, like these: http://imagej.net/Public_data_sets
Does anybody know any source of microscopic phase-images or does anybody have a set of complex images which could be used as a base?
I acquiered a small set using the quantitative Differential Phase Contrast Method provided by Lei Tian. These images are working quiet well :-)
Many thanks in advance!
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تجدها على النت يمكنك البحث
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Dear all,
Could you recommend any review paper (or book) comparing various downsampling methods applicable to volumetric data (preferrably, light microscopy or cell tomography data)?
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Some of publications of downsampling for volumetric data:1_ Optimal distribution_preserving downsampling for a large biomedical data.
2_ Downsampling method for medical datasets_Core.
https://Core.ac.Uk> download>pdf
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These are the structures from plant powders when observed under binocular microscope. Can anyone help me identify these
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Just try to add more data and also a scale bar. That will help.
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I'm working with enzyme immobilization on magnetic supports and I can't find specific protocols for characterization of these supports and enzymes after immobilization by microscopy.
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Dear all, the following documents give detailled experimental procedure in doing characterization of nanoparticles, and the information obtained via each technique. My Regards
10.1039/9781782621867-00001
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Hi,
I am planning to perform FTIR microscopy of brain tissues. I want to know that how slides can be prepared for this purpose. Can H and E stained slides be used or separate slides should be prepared on similar basis as for histology without staining. Later that slides can be stained with H and E stain after getting the peaks.
Please explain is it how we perform the FTIR microscopy and also that FTIR spectroscopy is better option to go for?
Best
Humna
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There is a lot of literature on this subject- but maybe you can start here:
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Hi everyone,
I am working on the microscopy simulation with Zernike aberration recently. Here I unfortunately fail to correctly add Zernike aberration to the simulated model or retrieve the aberration through Transport of Intensity Equation (TIE).
The details of this problem is as follows,
Here I have simulated a target with 0-1 intensity as File 'Sample' depicted. I also have simulated the Zernike Polynomials, for exmaple with the 7th order as File 'Zernike' depicted. I also assume that the numerical aperture (NA) of the microscopy is 0.45, and the wavelength of the illuminated light is 193nm. However, what makes me upset is that the Sample and Zernike are in different coordinates. The former is in Space Coordinate from -1200nm to 1200nm, and the latter is in normalized Pupil Coordinate from -1 to 1.
So, I wonder how to make them unity or indentical in calculation.
Thanks a lot!
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The following article might be useful
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I'm planning to modify a finite tube length compound microscope to allow the use of "aperture reduction phase contrast" and "aperture reduction darkfield" according to the following sources:
Piper, J. (2009) Abgeblendeter Phasenkontrast — Eine attraktive optische Variante zur Verbesserung von Phasenkontrastbeobachtungen. Mikrokosmos 98: 249-254. https://www.zobodat.at/pdf/Mikrokosmos_98_4_0001.pdf (in German).
The vague instructions state:
"In condenser aperture reduction phase contrast, the optical design of the condenser is modified so that the condenser aperture diaphragm is no longer projected into the objective´s back focal plane, but into a separate plane situated circa 0.5 – 2 cm below (plane A´ in fig. 5), i.e. into an intermediate position between the specimen plane (B) and the objective´s back focal plane (A). The field diaphragm is no longer projected into the specimen plane (B), but shifted down into a separate plane (B´), so that it will no longer act as a field diaphragm.
As a result of these modifications in the optical design, the illuminating light passing through the condenser annulus is no longer stopped when the condenser aperture diaphragm is closed. In this way, the condenser iris diaphragm can work in a similar manner to bright-field illumination, and the visible depth of field can be significantly enhanced by closing the condenser diaphragm. Moreover, the contrast of phase images can now be regulated by the user. The lower the condenser aperture, the higher the resulting contrast will be. Importantly, halo artifacts can be reduced in most cases when the aperture diaphragm is partially closed, and potential indistinctness caused by spherical or chromatic aberration can be mitigated."
The author combined finite 160 mm tube length objectives, a phase contrast condenser designed for finite microscopes, and an infinity-corrected microscope to get the desired results.
However, how would one accomplish this in the simplest way possible?
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Golshan Coleiny thank ýou for your reply. I assume you mean, for example, modification of the illuminator field lens to displace the conjugate aperture plane of the field diaphragm? Kind regards, Quincy
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I started my studies on histology and I realized most of well established text-books don't use scale bars in their images, except the books of the American-German Thieme Publishing Group. I think it is a lack of technical rigorosity, even though most of books treat the issue only on a qualitative point of view. So, what do you think about that? Should the publishers in that area change their concept to increase their accuracy?
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(only one point). maybe there is a difference to state how the image was made and how the scale could be, even though the scale bar is almost approximately at the same level each time. STM, (scanneling Tuning Microscope, false colour, the blacvk and white images, maybe the microscope taken with this 10x, objectives, and so on.All these give ideas on how the scale bars are.... even though I think it is ignored data.
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the question is related to microscopy
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Turn the scope on, remove the objective aperture, focus and center the beam (with the beam deflectors). As you over and under focus the beam, if the beam moves/shifts from the center, the condenser aperture is misaligned. Keep over and under focusing the beam as you manually adjust the X and Y of the condenser aperture. When there is no longer a beam shift as you over and under focus the centered beam, the condenser aperture is aligned.
B
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Hi, scientist community,
I am trying to improve the sample preparation for microscopy. The main problem is bubbles formation under coverslip when I put the coverslip on sample drop. Sample drop is in the center of spacer cavity. The spacer is an sticker for the microscope slide. Firstly I paste this sticker on the microscope slide, then I put the sample drop (I calculate the drop volume using cylinder volume equation based on sticker cavity properties) and finally I put the coverslip and I always see several bubbles. The problem is not only bubbles, also the mobility of this bubbles (they are not stable).
Could you help me to improve the samples preparation using spacers?.
Thanks in advance
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Thank you very much for all the answers¡¡¡
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I'm a microscopy assistant working in core facilities. I'm going to receive the nerve samples for TEM. The researcher perfused mice with 4% PFA and conducted immunohistochemistry labeling by using DAB staining. Then postfix with 2% PFA/2.5% glutaraldehyde in PBS for 3 hours. Wash the samples 3 times with PBS. The samples in PBS will arrive on Friday and process next week.
I'm curious if it is ok to leave the DAB stained samples in PBS at 4°C for 3 days? I will continue postfix with 2% OsO4 on Tuesday. How long can we leave the sample in PBS? Is the PBS going to affect the DAB stained over time? Any method/protocol suggested would be appreciated.
Best,
Khwannarin
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Since +/- the same Q was asked for via the MSA-Listserver I only would like to include my personal reply to the requester via the MSA listserver [cf.: http://microscopylistserver.com/ ] on 2/9/2021 without no reply still (perhaps dumped to the spam folder then):
"Hi,
dear Khwannarin Khemsom,
as of my understanding, (= dealing with transmission electron microscopical specimen preparation and processing) one usually would use 'phosphate buffer' instead of 'PBS (undefined)'…
So it might be useful and / or of benefit to the MSA_listserver community to define the „PBS“ you use (as you for sure know that there are many „qualities“ of PBS out there in the wild….protocols/recipes…with & without ions, different molarities (stock solutions, working solutions….etc…).
My best bet would be that storage in „ordinary“ phosphate =PO4 buffer (e.g. Sörensen/Sorensen) or also Na-Na (sodium-sodium-PO4 according to Millonig) (at 4°C) in the right molarity (i.e. usually around 0.1-0.13M) and correct pH (i.e., between 7.0 and 7.3) would be of no major harm than ‚incorrect applied‘ >PBS<…
Regards and
sincerely yours,
Wolfgang
PS: Admitting having done a lot of TEM on nerves…but not having been sent / received many specimens from „abroad“ or extramural for diagnostic examination ….nevertheless knowing about the artifacts improper handling (i.e. ‚handling‘ you don’t know about how it REALLY was performed) and/or extramural primary processing COULD induce major artifacts/damage to nerve tissue, especially after previous IHC-labeling (DAB used formerly e.g. for ‚pre-embedding‘ immunolocalization of markers)….
Hopefully you don’t feel wheedled…
Wish you well….and successful" (-End QUOTE-)
Retrospectively:
You might have a look into an old research article (uploaded in my Personal profile/publications, knowing there are tons of other specific and perhaps better research articles on the /your problem 'out in the wild') I was involved in performing the Electron Microscopy of specimens:
If performing only a 'small/short" query via Googleing | DAB precipitate | one might gain good insight of the process going on when using DAB [usuall recation HRP catalyzes the oxidation by hydrogen peroxide (H2O2) resulting in the generation of a brown / dark PRECIPITATE >>insoluble in water (hence most probably in 'physiologically adjusted' PBS or phosphate buffer), alcohol and other organic solvents (as used usually in the lab, e.g. isopropanol, xylene, etc.)....
NB (only trying to provide as much info as possible in this specific RG-thread) :
Another colleague answered there ('seconding' that answer 100%):
"DAB is a permanent stain and will not wash out with PBS, alcohol, or solvents such as xylene.
Paula Keene Pierce, BS, HTL(ASCP)....."(end quote)
(Paula's coordinates were included here but for reasons of Data Protection omitted here)
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My intention is to be able to see by LOM, the melt pools produced by the additive manufacturing process, and also the grain formation.
Any help or advice would be appreciated!
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Are there any tips for obtaining clear images (20x) of cell files in each zone of Arabidopsis roots?
I normally fix them first in 96% ethanol and mount them in chloral hydrate. However, due to the ethanol, the roots are wrinkled, starting at the elongation zone (the meristem is okay).
Reducing the timing of ethanol clearing does not alter this effect, nor does using a lower percentage of ethanol, as observed in my samples with GUS-staining which are conserved at 70%.
I could try increasing the percentage, but this will be time consuming to apply to each sample.
Does anyone have any other ideas?
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You may use visikol. Put the seedling in a slide, add few drops of visikol, put the cover slip and incubate for 10 minutes at 37C. Your root cells will be cleared like magic. We routinely use this in our lab. Previously we used to use conventional cell clearing method using acid-alkali and alcohol. You can find that method in Malay and Benfey Development paper on Lateral root. This method also works well but takes longer time. Visikol is little expensive but works like magic and shorten the processing time.
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I have grayscale images obtained from SHG microscopy for human cornea collagen bundles, and I have them as tiff stack images and their Czi format. I want to convert those 2D images into a 3D volume but I could not find any method that can be done using MATLAB, Python, or any other program?
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If you know the physical dimensions of your images and the images in the stack are properly aligned (consecutive), you can create a 3D volume in matlab, then write that volume as nifti (normally for neuroimaging, but should do the trick). There are many tools that can work with nifti, perform 3D volume rendering etc., such as 3D slicer. It is just a representation, the important thing is to have the mapping between physical and image coordinates.
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Hi,
I wanted to purchase indocyanine green from the sigma-Aldrich company they provided two types. I'd like to ask you what is the difference between these two phenotypes ( polymethine dye Vs. For Microscopy) of Indocyanince green (Cardio-green), which one best suited if to be used in a patient oral cavity? Does it make any difference if any of them is to be purchased, please?
Thanks
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Like Frank T. Edelmann said, there are usually versions appropriate for human/animal use and it doesn't look like either of the options you listed are. But they do have a USP grade option (product # 1340009) that meets or exceeds requirements of the United States Pharmacopeia (USP). This grade is acceptable for food, drug, or medicinal use.
Chemically, the two products you listed are the same. The difference between them might be purity. The version listed as "polymethine dye" has a listed dye content of ~85% but the "for microscopy" version doesn't specify purity.
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As I am going to create a raw microscopy image, I have to consider what possible noise might be present. To ensure I do not miss any important points in noise modeling, I am looking for a good literature as a reference.
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the following paper fits your needs:
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Did you ever use ruthenium red technical grade for TEM and if positive, did you see any difference in TEM images using ruthenium red technical grade vs microscopy grade?
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Dear Alexandra, in addition to the expert answers which have already been provided, please also have a look at the answers given to the following technical questions about ruthenium red which have been asked earlier on RG:
Questions related to Ruthenium Red
I would also like to suggest to you the following potentially useful literature references:
Ruthenium red and violet. I. Chemistry, purification, methods of use for electron microscopy and mechanism of action
and
Purity of ruthenium red used in pharmacological research
Unfortunately these two paper have not been posted as public full texts on RG. Perhaps you can access them though your institution. The authors of the second article both have RG profiles. Thus it might be worth a try to request the full text of the paper dierctly from one of the authors via RG.
Good luck with your work and best wishes!
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Hello,
I have a question regarding the microscpoy development.
In general, after the inital idea, do you have an idea on how long it took to build and develop until achieving an actual image.
I was searching for solid examples for microscopes like, spinning disc/ligh-sheet. But i couldnt find regarding the developmental process.
Thank you for your time
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It depends entirely on how complicated your system will be and on if the technology you need is available, or if you must develop the technology yourself. I designed and built a moire’ light microscope in about 1 week. But it took about 1 month to develop the software needed to do the actual analysis via computer-based image analysis. The microscope was a first of its kind and very limited to analyzing micro layered materials (that is, layer thicknesses across the entire sample in cross-section).
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which software is easy and best to use for SEM, scanning electron microscopy images analysis ..
such as for stomata area. trichome length. trichome tickness. fruit peel different layers.
etc
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It is better to use ImageJ.
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I need to perform some Scanning capacitance microscopy (SCM) & Spreading resistance profiling (SRP) on some samples. Thus I am looking for labs that offer this service.
thank you!
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Contact Aystorm Scientific Ltd ; We ll be delighted to help
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We are studying biofilm formation on differently coated 15mm diameter uPVC discs - these are opaque and may be white or black in colour depending on coating. The current setup includes forming biofilms on discs, rinsing to remove non-adherent cells, fixing each disc to a glass slide and then adding a drop of LIVE/DEAD BacLight mixture (Invitrogen L7012) to the centre of the disc. After incubation, the excess stain is rinsed off, a coverslip is placed on top of the disc and the edges of the coverslip fixed to the underlying slide with nail polish. After drying, the whole piece is placed in the inverted microscope coverslip side down. When imaging it is possible to see a hue of green or red depending on filter but not to resolve cells, and it is unclear whether this is just autofluorescence of the disc or coating. The opacity of the discs seems to block much of the light.
Thank you for your assistance and advice.
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Hi James,
The opacity shouldn't matter since you are illuminating as well as observing from the coverslip side, so neither excitation nor fluorescence need to propagate through the disk. We image biofilms on steel plates in a similar way. Is there water between the coverslip and the PVC disk?
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Due to a project about holographic microscopy, I'm gonna write a review proposal and for a part of that I need good literatures about Phase Retrieval from images using Fourier transform.
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I need s a testing plan for a stable anti-fade reagent for cell microscopy.
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I have never used and never heard ascorbic acid as antifade solution. I have never heard Trolox as antifade solution. From where you got these ideas?. Follow Ashten's suggestions, that will work
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I am wondering if there is a type of microscopy technique that people use in the field of insect physiology that allows muscle to be seen in living or prepared specimens.
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Hi Tristan -- I'm taking a look into this -- I bet you already found some answers on your own since a year has passed, but I'll look into it right now out of curiosity.
The best I could find from a quick search was a mice study here ( ) -- I particularly liked Figure 1 here -- and I am taking a look at Reference # 26, 27 ("A few studies previously reported in vivo microscopy of skeletal muscle" [I know you are not looking at skeletal muscle per se of course, since we are talking about insect muscles.]) . . .
Summarizing possible methods for what I think you are looking for:
From the 1st paper I linked: "Multilodal microscopy combining SHG and fluorescence detection" where SHG stands for second harmonic generation.
The methods from the three papers here seems to be called "intravital imagining of muscles" as a whole.
From the third paper (Ref 27) -- it mentions confocal, intravital, and superresolution microscopy . . .
The second paper (Ref 26) mentions intravital microscopy...
A quick search yielded this result for intravital microscopy:
Interestingly, for insects, when I tried to find "intravital microscopy and insects" -- I found this instead: https://www.microphotonics.com/x-ray-microscopic-inspection-of-insect-flight-muscles/ --the title is a bit strange -- but specifically, I am looking at the Micro-CT imaging part, as it can allow you to visualize the internal and external features of your insect without having to kill it. You may want to anesthetize the insects like what Dr. Ellis does with her flies, perhaps.
I think that can be a good start for now... I know this is an old post but this was an interesting and useful question nonetheless.
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I have been trying to take images of my cells using 40X lens. I focus my cells first with the 4X then 20X lenses; but whenever I switch to the 40X I can't see the cells, not even out of focus. All I see is blurred image of the plate. I tried zooming in till the lens almost touched the plate but to no use.
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Dear Ayman
Check that the condenser iris aperture is set to the objective numeric aperture of 40x. In another case, I suggest checking that you are using the proper technique. Check if using phase contrast or DIC is no longer suitable for visualizing your cells.
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I am looking for the books or best research articles for the fret-film analysis using microscopy.
To unravel the changes in host proteins after the mycobacterial lipids treatment.
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The FRET efficiency can be calculated by subtracting the donor intensity in the presence of the acceptor from its intensity after photobleaching the acceptor, and then normalizing this value to the donor intensity after bleaching.
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Hi all,
I would like to fluorescently label mRNA that we will be delivering to MoDCs. I'm going to perform microscopy (Axios Imager D2 if that's relevant) to see localization of the specific mRNA within the cells. This mRNA is from a vendor so I can't transcribe it myself. Any thoughts?
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It doesn't sound like a good idea to use a fluorophore-labeled mRNA for imaging purposes inside cells. Instead, you would try a hybridization probe against your mRNA with their related amplifiers to make it visible. Otherwise, labeled single mRNA molecules will likely be stay below the signal detection threshold under a microscope even with 63 or 100X objectives. I have been using the RNAscope system to visualize endogenous mRNAs at single-cell resolution, which I think what you need. In your case, you may request your vendor to use synonymous codons for certain amino acids if need in order to avoid cross-labeling endogenous mRNA. But sample processing is quite tedious, though could be very accurate.
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We find these structures in the urine filtrates of humans/canines exhibiting a similar host of symptoms, of unknown etiology. They can be highlighted using a variety of stains and are along the scale of 250-500 um. We greatly appreciate any thoughts or suggestions you might have as to what may be creating these structures, or a more specific name/identify for them.
microscopy
cytology
infection
bacterial nets
fungal nets
fruiting body
myxamoebae
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Pics do not look like the typical urine cast
maybe artifacts
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I am trying to convert some cif files to .xyz file format for TEM simulation with QSTEM. All cif files obtained via Findit2009 and I followed the guideline: https://www.physics.hu-berlin.de/en/sem/software/software_convert_cfg
However, only the Li2MnO3 cif files can not be converted. I tried three Li2MnO3 and did not succeeded, while other files (such LiMn2O4 ) do not have this problem.
I doubt that whether there are some errors for those Li2MnO3 files and want to know how to solve it.
I attached the cif files and the screenshot of the converting error (the jpg.)
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Sorry for posing another answer. This answer may help other researchers who are looking for the same information. I have created cfg file using MATLAB, as mention before. If you want to convert cfg file to xyz, you can use the building model tool that comes with QSTEM. Best of luck
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When should I use Western blot for appotosis study, instead of FACS and Immunofluorescence by microscopy..?
Thanking you..
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Thanks for your kind contribution for most valuable discussion @Malcolm Nobre and Archana Chaudhary.
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I am doing research on microplastics in fresh water and for initial screening i have to do microscopy. I have dissecting microscope to analyse. i want to ask do i need any pretreatment for fresh water i-e treatment with H2O2 or density separation? since my sample is almost clear.
secondly do i need to use nile red dye for simple microscopy under dissecting microscope ?
and last what filter is best to filter my sample? I have filters which are black in color pore size 0.2micrometer . pore size is fine but have any one used black filters ??
Please give suggestions i really need help. recommend me relevant literature if u have.
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Dear Asma,
This is a really good question. There are a number of methodologies available to separate plastics in freshwater, ranging from a simple salt solution (which I've used) to far more expensive methods requiring special liquids. The biggest issues are 1) separating the size fractions (0.2 micron might be too small - you'll get everything including some clay), and 2) removing organic matter. Since your samples are mostly clear, you might get away without the organic matter removal step. Density separation can be done using a NaCl solution, which is easy to do and low cost, albeit with lower recovery rates, especially of HDPE plastics. Other methods are detailed in Prata et al. (2019). Your method will depend on the objective of your study - are you attempting just to see if microplastics are present or are you attempting to quantify the types and sizes of the microplastics? The answer to that question will determine the lengths you'll need to go. If you are attempting to quantify, removal of organic matter is a must and you'll need a different high density liquid, such as NaI solution. I'm attaching two papers that are really quite useful as they have really good methods.
Sincerely,
Brannon
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Hello!!
Could anyone point out to me an efficient protocol for microscopy of grapevine shoot tips?
Thank you so much
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I was wondering if someone could share a protocol to fix my plant tissue that has some GFP-tagged endophytic bacteria. Can I fix it with aldehydes? If so, how long I can presente the GFP bioluminescence?
Best regards,
João
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Hi Saulo,
Thank you for the response. I was concerning if the GFP fluorescence can be damage after the fixative method. Do you have any ideia about it.
Thanks
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I have lots of different environmental isolates and I need to determine the number of bacteria in my cultures with an Improved Neubauer Chamber. But most of the bacteria are too difficult to visualize with light microscopy. I read in some places that I can use methylene blue to make the bacteria more visible as it gives a bright/dark blue color to the cells.
The problem is that all the protocols I found use heat fixation, that I believe cannot be used in any Neubauer Chamber as it would potentially deforme the chamber changing its volume. So, I was looking for a way to stain bacteria in suspension with methylene blue. Is that possible?
Thanks
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Hello, dear Sandhya Jayasekara , well done!
Perhaps - or better 'for sure' - I forgot to mention the vital stains / dyes just to challenge the specialized community....(;-))...
Sometimes the 'old-ancient' - literature already has solutions and provide valuable information about techniques used today as a 'kit' with marginal knowledge of what happens 'in the box' .....and I second your appeal / proposal to perform some trial experiment beforehand... cool !
It might be also there are other dyes than MB to be considered....
also cf.
Article Erythrosin B: a versatile colorimetric and fluorescent vital...
[https_XX_://www.future-science.com/doi/10.2144/btn-2019-0066, html and pdf, respectively.
(or:Article Vital Staining of Bacteria by Sunset Yellow Pigment
https_XX_://www.exeley.com/polish_journal_of_microbiology/pdf/10.5604/17331331.1234999);
Article Development of a Vital Fluorescent Staining Method for Monit...
(https_XX_://www.ncbi.nlm.nih.gov/pmc/articles/PMC92329/);
Article The effects of vital dyes on living organisms with special r...
(https_XX_://link.springer.com/article/10.1007/BF01686508);
Article Confusion over live/dead stainings for the detection of vita...
(https_XX_://bmcoralhealth.biomedcentral.com/articles/10.1186/1472-6831-14-2)
and many more...
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I have more than 10k images (Phase contrast, 10X) of hMSCs and I need to count the cells. Right now, I am using ImagaeJ jo manually count them in a 'tick-tick' fashion which is very laborious and time-consuming. As there is not a very sharp contrast between the cells and the background, can a MATLAb code be written to ease my work?
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You might found your answer but I am posting python code here ---might help someone else.
import numpy as np
import cv2
import time
count = 0
def draw_flow(img, flow, step=16):
h, w = img.shape[:2]
y, x = np.mgrid[step/2:h:step, step/2:w:step].reshape(2,-1)
fx, fy = flow[y,x].T
lines = np.vstack([x, y, x+fx, y+fy]).T.reshape(-1, 2, 2)
lines = np.int32(lines + 0.5)
vis = cv2.cvtColor(img, cv2.COLOR_GRAY2BGR)
cv2.polylines(vis, lines, 0, (0, 255, 0))
for (x1, y1), (x2, y2) in lines:
cv2.circle(vis, (x1, y1), 1, (0, 255, 0), -1)
return vis
def draw_hsv(flow):
h, w = flow.shape[:2]
fx, fy = flow[:,:,0], flow[:,:,1]
ang = np.arctan2(fy, fx) + np.pi
v = np.sqrt(fx*fx+fy*fy)
hsv = np.zeros((h, w, 3), np.uint8)
hsv[...,0] = ang*(180/np.pi/2)
hsv[...,1] = 255
hsv[...,2] = np.minimum(v*4, 255)
bgr = cv2.cvtColor(hsv, cv2.COLOR_HSV2BGR)
return bgr
def warp_flow(img, flow):
h, w = flow.shape[:2]
flow = -flow
flow[:,:,0] += np.arange(w)
flow[:,:,1] += np.arange(h)[:,np.newaxis]
res = cv2.remap(img, flow, None, cv2.INTER_LINEAR)
return res
if __name__ == '__main__':
import sys
print help_message
try: fn = sys.argv[1]
except: fn = 0
cam = cv2.VideoCapture(fn)
ret, prev = cam.read()
prevgray = cv2.cvtColor(prev, cv2.COLOR_BGR2GRAY)
show_hsv = True
show_glitch = False
cur_glitch = prev.copy()
while True:
ret, img = cam.read()
vis = img.copy()
gray = cv2.cvtColor(img, cv2.COLOR_BGR2GRAY)
flow = cv2.calcOpticalFlowFarneback(prevgray, gray, 0.5, 5, 15, 3, 5, 1.1, cv2.OPTFLOW_FARNEBACK_GAUSSIAN)
prevgray = gray
cv2.imshow('flow', draw_flow(gray, flow))
if show_hsv:
gray1 = cv2.cvtColor(draw_hsv(flow), cv2.COLOR_BGR2GRAY)
thresh = cv2.threshold(gray1, 25, 255, cv2.THRESH_BINARY)[1]
thresh = cv2.dilate(thresh, None, iterations=2)
(cnts, _) = cv2.findContours(thresh.copy(), cv2.RETR_EXTERNAL,cv2.CHAIN_APPROX_SIMPLE)
# loop over the contours
for c in cnts:
# if the contour is too small, ignore it
(x, y, w, h) = cv2.boundingRect(c)
if w > 100 and h > 100 and w < 900 and h < 680:
cv2.rectangle(vis, (x, y), (x + w, y + h), (0, 255, 0), 4)
cv2.putText(vis,str(time.time()), (x,y), cv2.FONT_HERSHEY_SIMPLEX, 1, (0,0,255),1)
cv2.imshow('Image', vis)
if show_glitch:
cur_glitch = warp_flow(cur_glitch, flow)
cv2.imshow('glitch', cur_glitch)
ch = 0xFF & cv2.waitKey(5)
if ch == 27:
break
if ch == ord('1'):
show_hsv = not show_hsv
print 'HSV flow visualization is', ['off', 'on'][show_hsv]
if ch == ord('2'):
show_glitch = not show_glitch
if show_glitch:
cur_glitch = img.copy()
print 'glitch is', ['off', 'on'][show_glitch]
cv2.destroyAllWindows()
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Does anyone have any experience/reviews of the EVOS XL Core microscope? 
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Does anyone know how to insert a scale bar on an image using evos xl core? This option is not shown to me on the screen. I’m not sure if it’s not updated or if I have to set this somewhere.
Any ideas?
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Does anyone have a protocol for the usage of Mitotracker green to measure mitochondrial mass by a fluorometric assay? I was searching in literature and only find protocols for flow cytometry or microscopy, does anyone knows if it is possible to do it by other methods using this dye? It was to confirm some western blot results.
Thank you in advance
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You can use fluorescent plate reader as well. Label your cells with MitoTracker green following usual protocol and wash the cells extensively with pre-warmed media. Read your plate at ex - 488nm and em - 530nm. Be careful that this protocol is not very accurate compared to flow.
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Does anyone have any good tips on how to distinguish between hippocampal pyramidal and basket cells?
I am studying the morphology (dendritic branching and spine counts) of hippocampal neurons in golgi-stained mouse tissue. I was initially looking at pyramidal neurons, but my PI suggested I could look at basket cells too. However, when I looked at the morphology of basket cells in the literature/on neuromorpho,org, they looked similar to some of the neurons I had been labelling as pyramidal neurons.
I know that typically pyramidal neurons have a singular long apical dendrite that extends for some distance before other dendrites branch off it, while basket cells have multiple branches from at or close to the soma. Also that neurons with soma clearly within the pyramidal layer are more likely to be pyramidal neurons, but neither of these criteria feel clear cut enough for me to be confident. Sadly the basket axon doesn't seem to be labelled in any slices as far as I can tell.
I've attached a few images of cells I have included. They are collapsed z-stacks of about 20-40 um in z-axis. If anyone could give their opinion on what cell type they are and why, that would be very helpful.
Thanks in advance!
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Hi Craig,
Trying to differentiate between basket cell and pyramidal in golgi-stained sections may not be the best. Also, golgi works best for spine analysis in my opinion and not easy for dendritic analysis either and axons usually don't get stained with golgi (I never saw in my sections).
Having said that, the first image is a pyramidal for me, the second one likely, the third seems so but it's difficult to say. Also in your images, it's not clear which layer are you in. There seems to be less. The staining of adjacent cells in the hippocampal layers make it easier to categorize the neurons as pyramidal cells as they look quite alike with their somas packed next to each other and in similar orientations. But basket cells, as you already mentioned are not usually arranged in the same axis and orientation, if you can use that fact to differentiate between the two (maybe? I am not sure).
Here is a paper from our lab with some nice golgi images from CA1,CA3. Idk if it may help
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Hi everyone!
As I wrote in the question, I want to know if there are protocols to test spore vitality in mosses
I read about hemocytometer test, but are there some staining techniques that I could apply?
Many thanks!
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Updates
Germination in liquid Knop medium of T. muralis spores after 3 weeks incubation
It is not axenic just a try, now I'm starting to deal with contamination
See some photos in the attachment
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I am a microscopy image analysis noobie.It'd be great to have a discussion on this question.We have multi channel fluorescence images of dendrites and other brain structures obtained before and after an experiment on mouse.In the images obtained after the experiment,in some channels,fluorescence from 2 different viruses are present.Is there a way to separate the two fluorescence?
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I fully agree with Alexander Dräbenstedt that only mathematical postprocessing can give the solution to your problem. You can use a multichannel fast Fourier transform (FFT) preprocessing as well.
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I want to have a quantitative analysis of the excited molecules in 2Photon Excitation microscopy and compare it to the confocal one. Therefore if I can have the illuminated volume of the sample in these two techniques then I can approximate the excited number of molecules in that volume.
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A few things to remember:
- For the confocal microscopy, it is the wavelength of the light, the NA of the objective and the pinhole size determines the excitation volume in the Z space. The excitation light will excite a larger volume above and below the focal plane in the Z space, but the emission light is detected only from the confocal plane due to the pinhole blocking the out of focus light from being detected.
- When it comes to two-photon microscopy is its ability to restrict excitation to a tiny focal volume in thick samples. The objective focal point is the only space with a high enough photon density to ensure simultaneous presentation of two photons to the fluorophore. Effectively, this means there is no out of focus emission light and any light at the emission wavelength must have come from that single spot.
To learn more about the differences:
- Two-Photon Microscopy by Kurt Thorn (https://youtu.be/CZifB2aQDDM)
Good luck!
@Sathya Srinivasan
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I am having trouble with etching the heat treated Inconel 625 MIG weld sample. The heat treatment conditions are 980C with 30 mins, 1 hour, 2 hour. I have tried with Glyceregia, Mixed Acid (HCl, HNO3, Acetic Acid). Samples are getting lightly etched at first. Higher Magnification image like 400x are not good enough with this kind of etching. Etching again, I am seeing so many scratch marks (photo attached). If I can even over-etch the sample I can get to see the images in SEM. But during over-etching these scratch marks are occurring.
Anyone know what to do?
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Hi Jitender Kumar Chaurasia , please check this paper out -
You can find the etchant and way to etch in the experimental section
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I’m working on an image processing project with aim of developing A method to differentiate between healthy and diseased cornea. I have images taken from SHG microscopy for the collagen bundles in the human corne. I have used GLCM and I obtained the correlation, energy, homogeneity and contrast of each image but the results are not good enough to use them as a solid evidence to classify the images. please can anyone suggest for me any other method that can be used to classify collagen bundles image that can be done witthin a short time or a method to enhance my glcm results.
Thank you in advance
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Also, which parameters that can be known by using it, example diameter, feret diameter, perimeter of particles of the sample?
I want to study variation of salt hydrate grains on exposure to varying temperature and humidity. Will it possible with HSM?
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Dear Aastha Aastha,
I have attached the ASTM standard of hot stage microscopy(HSM). Every thing has been explained in details. I hope it would be useful.
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Hi dears
I calculated d- spacing my tem microscopy image but I don't know how can find miler index
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