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.
Questions related to Microscopy
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?
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?
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
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.
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!
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?
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?
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 =)
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.
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?
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.
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?
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?
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?
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.
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!
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.
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.
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
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?
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?
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!
Could you recommend any review paper (or book) comparing various downsampling methods applicable to volumetric data (preferrably, light microscopy or cell tomography data)?
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.
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?
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!
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?
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?
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
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.
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!
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?
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?
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?
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.
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?
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
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.
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.
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.
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.
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.
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.
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.
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?
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.
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.)
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.
Could anyone point out to me an efficient protocol for microscopy of grapevine shoot tips?
Thank you so much
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?
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?
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?
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
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!
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?
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?
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.
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?
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
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?