Microscopy - Science method

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

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).

Resin/plastic is used for electron microscopic studies and to examine hard tissue, such as bones.

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.

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.

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 .

I agree with Annemarie Honegger.

Dear Meagan Doppegieter ,

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

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.

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.

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.

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.

Zeiss products good in microscopy, I have used one stereo

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.

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 :)

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:

Methyl alcohol is the best

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.

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.

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

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

Mahyar Radak

These links might be helpful, have a look;

1. https://www.mathworks.com/videos/introduction-to-matlab-with-image-processing-toolbox-90409.html

2. https://www.mathworks.com/matlabcentral/answers/169745-how-to-generate-a-digital-hologram

Dear Neda Karami

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.

تجدها على النت يمكنك البحث

Some of publications of downsampling for volumetric data:1_ Optimal distribution_preserving downsampling for a large biomedical data.

https://www.ncbi.nib.gov>Pmc

2_ Downsampling method for medical datasets_Core.

https://Core.ac.Uk> download>pdf

Just try to add more data and also a scale bar. That will help.

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

There is a lot of literature on this subject- but maybe you can start here:

The following article might be useful

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

(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.

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

Thank you very much for all the answers¡¡¡

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,

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<…

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:

,

or also: and /or:

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%):

Paula Keene Pierce, BS, HTL(ASCP)....."(end quote)

(Paula's coordinates were included here but for reasons of Data Protection omitted here)

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.

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.

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.

the following paper fits your needs:

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:

I would also like to suggest to you the following potentially useful literature references:

and

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!

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).

It is better to use ImageJ.

Contact Aystorm Scientific Ltd ; We ll be delighted to help

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?

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

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.]) . . .

Ref 26 article:

Ref 27 article:

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.

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.

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.

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.

Pics do not look like the typical urine cast

maybe artifacts

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

Thanks for your kind contribution for most valuable discussion @Malcolm Nobre and Archana Chaudhary.

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

Please find the protocol for microscopy of grapevine shoot tips attached.

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

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...

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()

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?

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.

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

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

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.

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:

- https://www.microscopyu.com/techniques/multi-photon/multiphoton-microscopy

- Two-Photon Microscopy by Kurt Thorn (https://youtu.be/CZifB2aQDDM)

Good luck!

@Sathya Srinivasan

Hi Jitender Kumar Chaurasia , please check this paper out -

You can find the etchant and way to etch in the experimental section

Other then the conventional statistical approaches for textures analysis , you may also follow https://www.nature.com/articles/s41598-020-67632-z

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.