Confocal Fluorescence Microscopy - Science method
Confocal ﬂuorescence microscopy is a microscopic technique that provides true three-dimensional (3D) optical resolution. This technique has gained popularity in the scientific and industrial communities and typical applications are in life sciences, semiconductor inspection and materials science.
Questions related to Confocal Fluorescence Microscopy
Does anybody know of a live membrane dye which works well with tissues and isn't internalised or pumped out of the cells in a short time frame? I'm looking to perform 1 or 2 hour imaging experiments on live vessels (ex vivo), however most of the live membrane dyes my lab has previous experience with are only really suitable for very short-term experiments. I should also add that due to the nature of the experiments I plan to carry out, using probenicid to retain the dyes for longer won't be an option.
Any advice would be greatly appreciated! Thanks!
I am trying to immunostain whole (micro)tissue samples, and I think there is an antibody penetration issue. Would longer permeabilization procedures or harsher permeabilization agents help with this?
Thanks for sharing your experiences!
Could you please recommend DAPI concentration for nuclei staining in mesenchymal stem cells during chondrogenic differentiation?
I have used 1 μg/ml DAPI for 6 min, then as usual PBS washing. but I got fuzzy colour and lots of unspecific binding (but still fuzzy).
If that matters - the brief protocol:
- PBS washing 3x
- permeabilization 0.2% Triton X-100/PBS, 4 min, PBS washing 3x
- blocking 0,2% BSA/PBS 1h, PBS washing 3x
- staining of collagen I, II and X (each with a pair of antibodies, with 3x washing in PBS in between steps).
I am going to analyze differentiated cardiomyocytes (CM) via confocal microscopy. However, I struggle to attach CM properly to the glass slide/plate. Although I coated the glass slides with LM-E8, the CM could not attach firmly and started to detach after adding 4% PFA to cell staining.
Does any appropriate coating for CM culture over the glass?
I appreciate any recommendations and comments.
I'm just starting to teach myself how to make macros to process confocal images in ImageJ (FIJI). I've managed to make a macro that does most of my processing using the record macro function (it has already saved me an enormous amount of time!), however I'd like to add brightness and contrast adjustment to the processing.
My current process involves:
1. Splitting the multi-channel images
2. Applying grey LUT to all
3. Saving all in a temporary folder
4. Creating a merged image of the channels is various colours
5. Creating a panel with split and merged images
6. Adding a scale bar
7. Saving and closing all windows
However, when I look at the recorded code produced by the brightness and contrast adjustment dialogue, I get code which doesn't seem right to me:
I've also tried the following code, however it automatically maximises the contrast, where I'd like to use the brightness and contrast adjustment to reduce background in some channels.
run("Enhance Contrast", "saturated=0.35");
Is there anyone else using macros for brightness and contrast adjustment in ImageJ?
I am facing a very intriguing phenomenon. My immunofluorescence and confocal microscopy analyses showed a nucleus that is positive for MnSOD (which should be present only within mitochondria). Please, find attached representative images of cells stained for either nucleus or MnSOD positivity, along with the merged image.
Of note, it should be highlighted that the anti-SOD2 primary antiobody we used is extremely specific and that Western immunoblots showed only the specific band.
Did anyone else find MnSOD (SOD2) in the nucleus in some experimental or physiological condition?
Thanks a lot in advance.
- 461.52 KBSHSy5y ELF SOD1 e SOD2 5 e 10 gg 21.12.2016_Controllo 5 gg SOD2 zoom 2 1 Series011_z0.tif
- 208.56 KBSHSy5y ELF SOD1 e SOD2 5 e 10 gg 21.12.2016_Controllo 5 gg SOD2 zoom 2 1 Series011_z0_ch00.tif
- 270.31 KBSHSy5y ELF SOD1 e SOD2 5 e 10 gg 21.12.2016_Controllo 5 gg SOD2 zoom 2 1 Series011_z0_ch01.tif
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 looking for a nuclear marker, like a transcription factor, expressed in human enteric neurons.
I would like to characterize my human iPSC-derived culture with immunofluorescence stainings. The problem is that my neural progenitors give rise to enteric neurons and enteric glia (GFAP+), so I cannot use Sox10 throughout the whole maturation. So far I haven't found anything this specific.
Does any of you have any suggestions about a good antibody, or a marker that would be useful in this case?
Thanks a lot in advance.
I am currently using Jasplakinolide to treat my Be2-C cells and I want to measure cell viability. I want to fix the cells in PFA and mount them with Vectasheild. However, I have seen a lot of different ways to use PI with PFA fixation, and I am a little unsure the best way to do it. I am thinking I will first treat the cells with PI by adding it directly to the media in the incubator for 1 hour and then fix with PFA, stain with DAPI, and mount? Any advice/suggestions?
Also, has anyone used this and also stained for phalloidin? I am worried there will be too much cross talk to use both.
Thank you in advanced :)
I have a FRET system between the inner and outer nuclear membrane, and I was wondering if I stained with DAPI or any other fluorescent antibody if that would mess up my FRET readout? As of now I am using acceptor photo bleach and I am working with neuroblastoma cells (Be2-c). I can do either fixed or live cell imaging, depending on which would work best if this even possible.
I made a script to try to convert 4 colour nd2 files (not stacks) to run in the process> batch function of Fiji (see https://www.youtube.com/watch?v=fU104OU3Kk4 at 3:28) to create single tiffs, of all colours, merged. When I check the results in Fiji I get only the DAPI channel from the script below. However, if I run the macro in Fiji itself without the batch command, just on a single open nd2 file, I am left with the file that I want! So I am not sure why it is only saving the DAPI. Does the batch command save the earliest tiff that it sees and not let the macro finish? I am an inexperienced Fiji user so I am not sure what I am doing wrong.
title = getTitle();
one = "C1-" + title;
two = "C2-" + title;
three = "C3-" + title;
four = "C4-" + title;
run("Merge Channels...", "c1=["+one+"] c2=["+two+"] c3=["+three+"] c4=["+four+"] create");
I have isolated mitochondria from tissue. I would like to check them by fluorescence imaging. Can I use Mitored dye for these isolated mitochondria?
I want to detect intracellular bacteria in urine by confocal fluorescence microscopy
I would like to build a lateral projection of a z-stack with the correct linear size ratio for the final illustration, how can I change the pixels size ratio in ImageJ?
I'm observing a small tumor with confocal. I imaged the expression of a fluorescent protein, which is supposed to express everywhere. However, I can see some vague DAPI signals, where I can't observe the fluorescent protein. Is this normal? If so, wouldn't the images that I obtained for a single layer be always interfered with vicinity layers?
I am wondering why my hepg2 cells nucleus looks fuzzy and unclear under the confocal?
Is this mycoplasma contamination?
I used Hoechst staining, 1:200 dilute with 3%BSA.
I am working with cells labelled with Mcherry (membrane) and EGFP (cytoplasm). Following my experiment, I want to quantify the green signal which relocated to the membrane. How can I extract and quantify in Fiji the amount of green signal on the membrane (e.g. where the red signal is)?
Thanks in advance for all your answers and suggestions!
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 z-stack confocal images of whole-mount tissue IF staining, marking the tight junction of the cell. And I would like to do 3D reconstruction of the cells to measure cell circularity and area using Imaris.
However, since it's challenging to mount my tissue perfectly flat, my z-stack images are slanted.
This makes the cell reconstruction challenging since the cell membrane signal for a certain cell is spread across multiple z slices. I attached few screenshots to demonstrate.
I'm wondering if there is a way to rotate my z-stack image and reassign the xy reference angle to computationally make the image flatter?
I am using a BD pathway imager. When the spinning disk gets in the light path I get very blurry images( see attachment). Does anybody know how to fix this issue. First image is with regular fluorescence image. The second is with confocal at the same focal plane.
I am working on amplifying a signal for confocal imaging using the Labelled Streptavidin/Avidin Biotin (LSAB) technique and ran into a surprising outcome: No staining. This is odd as I've employed this stain previously using the traditional indirect method and see decent specific signaling so I thought by amplifying the signal I'd get better, not worse, staining. Below I'll leave the basic outline as to my design (barring washes in between and counterstains).
- Incubate mouse anti-antigen primary antibody at 4 degree overnight (1:100; used at same concentration as in indirect method staining)
- Incubate biotinylated goat anti-mouse secondary at RT for 90 mins (1:150)
- Incubate Cy3-conjugated streptavidin for 90 mins (1:1000)
Any tips as to how to optimize this protocol and get it to function would be greatly appreciated!
When performing an indirect immunofluorescence experiment, it is not favorable to use a primary antibody raised in the same host as the tissue you will be staining (ie. mouse anti-x on mouse tissue, rat anti-x on rat tissue) due to the increased background you will get from the secondary antibodies binding with endogenous IgGs. Logically, it follows that if I use a fluorophore conjugated primary antibody for direct detection, this issue will not arise despite the antibody being raised in the same host as it is reactive for because there are no secondary antibodies involved. Can anyone verify that my assumption is true? If not, please share with me why my rationale was incorrect. Thanks!
I am working with FFPE sections of mice ileum and am trying to image changes in the mitochondrial ROS production between a treatment and control group. I'm not sure if there are any stains or antibodies that can specifically target mtROS once the tissue is fixed and I have seen another researcher propose using MitoSox Red on tissue prior to fixation to get an assessment of mitochondrial ROS in the live animal. Has anyone else tried this successfully?
We also dabbled using a pimonidazole hypoxyprobe as a measure of total ROS, however the staining in the ileum and colon differ for physiological reasons. If anyone has some ideas as to how to better optimize this stain or an alternative method of detection that works well with ileal tissue, please let me know!
If a sample has both GFP and RFP expression, and GFP is activated by a laser, then will its emission light be absorbed by RFP and cause RFP to shine? Another toy example would be, will the emission light of Alexa Fluor 555 become the excitation light of Alexa Fluor 647?
If not, why? If so, how could this be resolved during confocal imaging of dual-color? I don't think this is FRET, as FRET doesn't involve an actual photon being emitted..
Thank you so much!
I'm having problems with the Recombinant Anti-MAP2 antibody from Abcam (Recombinant Anti-MAP2 antibody [EPR19691] (ab183830)) antibody on my frozen, 4%PFA perfused, mice brain tissue.
I was wondering if someone could elaborate better the note from the application part of the data sheet; Antigen retrieval: Heated citrate solution (10mM citrate PH 6.0 + 0.05% Tween-20).
To my knowledge AR is not used with frozen tissue, as it is to rough, and can damage the tissue, while also causing tissue detachment from the glass!?
Does someone have any experience with AR and frozen tissue?
I did try using 1/100 dilution, 0,2% triton X-100 and 4%PFA fixation as described in the images part of the datasheet: Immunohistochemical analysis of 4% paraformaldehyde-fixed, 0.2% Triton X-100 permeabilized frozen Mouse Cortex tissue labeling MAP2 with ab183830 at 1/100 dilution.
However I did not observed cytoplasmic staining of my neurons (image as uploaded for reference)!!
My protocol was the following:
4%PFA fixation, 15 min RT
10 min PBS+0.2% Triton x-100
Blocking solution (3%BSA, Without Triton x-100, 1h)
Anti-MAP2 1:100 dilution in 1%BSA (try both overnight and 2h RT)
AF-647 1:500, 30-45 min RT
3xPBS, 5 min
DAPI, 5 min
3x PBS, 5 min
dH2O, 5 min
Antifade and coverslip.
The antibody works excellent in IHC-P.
Thanks for the help,
I am trying to characterize the microstructure (localization of carbohydrates, fats, and proteins) of coconut milk powder using confocal microscopy. From the review of the available literature, I have found that fats and proteins can be non covalently labeled using fluorescent dyes. However, I am having trouble deciding the labeling protocol for the carbohydrate. Two common methods that I have seen in the papers are as follows:
1. Use antibodies or lectins conjugated with fluorophores to label carbohydrates
2. Covalent labeling of carbohydrates with FITC
If I use the first method, the signal from antibodies or lectins may interfere with the signal from protein present in the sample.
In the second method, the FITC (probe used for covalent labeling) also shows an affinity for proteins. So, I am afraid that the use of such a probe may also interfere with the protein signal (from the food sample). Is there any other probe available that I can use?
What would be the best way to label carbohydrates such that I can simultaneously image three components in the food mixture?
I did a Western Blot for a protein that should only be present in hematopoetic cells. The WB shows a very clear picture, present in k562(hematopoetic) and not present in HeLa and 293T. However when using the exact same antibody for IF on HeLa cells we do see clear signal from our protein of interest (see picture). Can anyone provide insights to how this is happening? in controls without primary/secondary antibody we see no signal.
Hi! I am about to use AGR2 protein in my antibody display system. I want to check wherther protein can bind to cell surface displayed antibody or not. I need suggestions on how to fluorescently label (with red dye) AGR2 protein in order to view binding and also if someone could suggest efficient and cheap method to label protein that would be really great. please give me some insight and useful tips!
I am a physicist, so I know the techn not the biological applications. The question is straightforward. I am looking for a diagnosis that rely on fluorescent based cell imaging where the intensity of the expressed marker is the key part of the diagnosis. Thank you !
Looking for recommendations on what to use for imaging spheroids and organoids using a confocal scope. Would it be feasible to carefully transfer spheroids to glass bottom dish with coverslip, or better to use glass bottom microplate? Or another method altogether? I am experienced in the formation of spheroids but not so much the characterization. Thanks in advance
I have dozens of confocal images to process in Fiji. It is neither practical nor a productive use of my time to do them all out by hand, so I want to use a macro. However, I am fairly new to research; I only started a few months ago, so I don't have much experience writing macros in Fiji. What I'm doing is taking an image split into 3 channels, applying the grayscale LUT, and z-projecting to max intensity. Then I manually adjust brightness/contrast depending on the quality, merge channels, and stack to RGB. I also add a scale bar, but that I adjust manually i.e. 10 microns, 20 microns, etc.
Does anybody know if such a macro exists (i.e. in the Fiji user guide) or how to write such a macro? I have made one by using the record function, but that turned out to be inefficient because I had to change the file name every time I ran the macro. is there a way to get around this? Also, how would I incorporate into the macro the pauses when I have to make manual adjustments?
I use Richardson-Lucy deconvolution algorithm (flowdec package) with custom generated PSF. And after several iterations, granularity is observed in the cytoplasm, but the protein must be distributed diffusely. Random noise fluctuations resulting from deconvolution become bright points in the cytoplasm.
How can I reduce noise influence and prevent occurrence of an artifacts? And/or could I implement regularization (Tikhonov regularization f. ex.) in flowdec package?
Basically I would like to visualize (either by confocal or electron microscopy) differences in transcriptional activity inside a tissue, to see if I have cells that are more active in gene transcription then others . I am using an entire organism, a marine invertebrate animal, not a cell culture.
For ex. a staining method that will show euchromatin (non-condensed DNA) activity in the nucleus?
Also, somebody mentioned to me using the bromouridine labeling (Bru) to asses changes in RNA synthesis. Did anyone try this method on an organism?
Thank you !
I have a protein (conjugated with FITC fluorophore) that may adsorb and cover the bacterial cell surface or be taken up into the cytoplasm.
Using confocal (fluorescence) microscopy, how can you tell the difference with any confidence? (LSM700)
The cells are rather small at 200 x 1200 nm.
I'm relatively new to microscopy imaging analysis so I'm seeking some help! I have z-stack images (.czi files) from zebrafish using a Zeiss LSM 880 confocal microscope at 40x water immersion objective. My advisor has suggested using the ZEN software to do a maximum intensity projection and then using orthogonal view. The images still look "messy" after conducting these steps in the ZEN Blue v3.1 software, so I'm wondering if you have any suggestions or protocols to analyze images. Ultimately, I would like to compare fluorescent intensities, myelin sheaths/olig, and/or internode length across my samples. (also- should I implement a deconvolution step?)
Thank you in advance!
I have a grayscale image of which the original RGB image was formed by combining Green and Red channels which represent different types of cells (live/dead). Splitting the channels in RGB mode is giving Red, Blue and Green to all the cells. I want to confine green to the live cells and red to the dead cells using ImageJ.
Dear fellow image enthusiasts,
I just recently started doing a lot of confocal fluorescence microscopy imaging and I am getting good images, but I still thought about increasing their quality. I immediately considered deconvolution methods. Only problem is, I have never used a deconvolution tool before.
Could anybody help me out with kind of a "quick start protocol"? I want to use imageJ and donloaded deconvolutionlab 2 (http://bigwww.epfl.ch/deconvolution/deconvolutionlab2/) as well as the PSF generator they recommend (http://bigwww.epfl.ch/algorithms/psfgenerator/).
Which method should I chose to generate a PFS which would work for deconvoluting my 2D image?
Which parameters do I have to know (both while taking the image and for generating a proper PSF)?
- I do know the refractive index if my immersion oil, do I also need to consider the refractive index of the NPG used for covering and/or of the glass cover slide?
- I also know, the NA
Which method should I chose for decovoluting the 2D image?
I made a couple of test runs, but the output looked much worse than the input and particularly proper colour information was lost.
- 165.35 KBC1-Composite.png
- 154.22 KBFinal Display of RIF1.png
- 368.80 KBFinal Display of RIF Composite (RGB).png
- 102.21 KBC2-Composite.png
- 315.68 KBComposite (RGB).png
- 155.37 KBFinal Display of RIF.png
I am working on Symbiodinium cells (symbiotic dinoflagellates) and am trying to calculate the mitotic index. All the papers I have referred to use DAPI. I want to know if there are other stains or techniques that an use to distinguish between the doublets and just single cells. Is the resolution of DAPI good enough for me to see the mitotic spindle?
I have localised a protein in Endoplasmic reticulum of Saccharomyces cerevisiae. I have tagged this protein with HA tag. Now i want to label this protein with anti-HA fluorescent antibody for FACS and microscopy purpose, If anyone can suggest me a protocol for this then it will be a great help.
I have been imaging many z stacks of Drosophila mid/hindgut tumors using a leica confocal microscope. I'm wondering what the best way to quantify these tumors would be. They are GFP+.
I know both leica and image J have quantification programs. Any preference? Any recommendations would be greatly appreciated!
I need to image cyanobacteria using fluorescent microscopy, I have easily done this before with good results using the fluorescence of chlorophyl on a traditional fluorescent microscope. However, when I attempted to do this on a confocal system I had very poor results when imaging Cy5.5 along side the chlorophyl.
What excitation and emission wavelengths should I use for chlorophyl, specifically for cyanobacteria?
Should I use a different second dye like NBD or rhodamine instead of Cy5.5?
I'm running a study in which I will inject some nanoparticles into mice and then section the lymph node and visualize it with confocal. Does anyone have the protocal for the lymph node sectioning and staining? Thanks.
I'm working with a Leica SP8 confocal microscope,
and I'm trying to analyze the sum intensity of z-stacks (16-bit).
For particle analysis I'm making a max intensity image first which later I redirect its ROIs to the SUM.
A) in certain images I'm using the "make binary" and the output image is kind of clean B&W, in other images (from the same project and settings) I just get a cloud of pixels (I guess it's over saturation) among my cells.
B) in order to maintain a proper workflow I'm trying to batch analyze my files (8-bit->8-bit->threshold->analyze particles etc. etc.).
I would like to analysis the S. aureus biofilms to know the effect of certain quorum sensing inhibitor on ultra structure. Could someone elaborate the method to prepare sample for TEM analysis (Fixation, dehydration, drying, staining, embedding, ultra sections etc)? Many thanks for your valuable time and help
We are attempting to study actin/cytoskeletal function under different conditions in live cells. Does anyone know of an existing cell line that stably produces GFP-actin for purchase? I know that this can also be accomplished by transducing cells with viruses if you have any products in particular you recommend.
We are also ok with accomplishing this by labeling. I have tried BacMam and SiR-actin with fairly low efficacy in HeLa cells just as a trial (ideally something that works with easy to transfect and primary cells preferred). Considering trying LifeAct products but not sure if they are any better. Any recommendations on products and exactly how you applied them (number of cells, volume, inc time, concentration, etc) would be highly appreciated.
Can we use Tween 20 instead of triton X in 0.1% PBT-T(I assume T stands for triton X only) for immunofluorescence ?
Also, can tween 20 be replaced by tryton in IP wash buffer?
I found that the signals for my dye are weak in tissue sample, what is the maximum limit to which I can increase the detector gain and laser power percent for my dye?
I am going to mesure the mitochondrial calcium uptake with Rhod-2AM in permeabilized neonatal rat cardiomyocites, but the fluorescence signal is altered due to cell contraction, generating an artifact signal.
I read about 2 pharmacological agents to disrupt the excitation-contraction coupling: -blebbistatin and 2,3-butanodione monoxime. It seems to be that both pharmacological agents are not frequently used, and have been descibed different work concentrations.
What is the best option to inhibits the excitation-contraction coupling? and what is the work concentration?.
The literature is plenty of examples of immunostaining in live cells, but is hard to find the same in live organotypic culture. Does anyone have experience with this? Is there (a priori) any caveat about this technique?
By looking excitation spectra I would say no (~5% excitation), but its maybe enough in practice ? Has anyone already tried ?
Thanks, have a good day.
I would like to observe microgel particles inside fibres using UV microscope, thus I came acrross some difuculties when choosing the appropriate UV dye. The microscop can emit light at 405 nm, 488 nm, 555 nm and 633 nm. The dye has to be soluble in water, as other solvents like methanol and DMSO damage the fibres.
I've measured the diffusion coefficient of a fluorescently labeled glycosidase interacting with its substrate (a polysaccharide) by using fluorescence correlation spectroscopy (FCS). An example can be seen in the attached figure. The substrate was in its solubilized form and in saturating concentration (5 mg/ml). After FCS curve fitting, I found that 75% of the species corresponded to the free enzyme (Rh = 2 nm), while 25% were a much bigger particle (Rh = 37 nm). This value matches very well to the Rg of the polysaccharide (31 nm) according to the manufacturer. The effect of viscosity is already taken into account. However, since many enzyme molecules can be simultaneously associated to the carbohydrate, these 25% may not correspond to the actual proportion of enzymes in the bound state.
If my reasoning is correct, my question is: How to determine the average number of enzymes associated to the carbohydrate? I suppose I should analyze the photon counts over time, but I'm not sure how to do that.
Thank you in advance!
I have a set of confocal microscopy images. I am finding little difficult to conclude from the data, so need help from experts on this. I will upload the images as someone is interested.
Did anyone observe the suppression of low fluorescence signals from one organelle due to high signals from other organelles?
Hi, I want to quantify total tau, phospho tau and tubulin in the axon and dendrite of hippocampal and cortical neurons by Confocal microscopy analysis. I'll appreciate if you could suggest me the method to distinguish axon from dendrites, and the method to quantify them using imageJ software.
I have attached the image file/picture I scanned using microscope (red channel: total tau, green channel: tubulin, yellow channel: phospho tau). The image is focused on neurites.
I want to microscopic studies of cultured unicellular algal cells. I want to know which dye is most widely used for nucleus staining for algae.
I'm studying the gelation process of certain bio-polymers. When I form the final gels and cut it into small pieces for observation under the Confocal microscope. This process seems to hamper my gel structure. Is there any other process wherein I could form the gels in accessories that could be directly used for observation under the microscope. Please bear in mind that I have to heat the gels to induce gelation, so the material needs to resistant to heat. I have read of quart glass dishes can anyone suggest anything else?
Leica and Abberior say: ¨Please do not use Vectashield, Vectashield Hard set or (other) embedding media containing p-phenylenediamine as antifading reagent." (I do not understand why, but I have no choice to trust them)
Prolong Gold transforms the cells (10 - 12 um of the thickness) in 2 um pancake
The list of allies of a microscopist becomes short ...
I am currently writing my first master project. I have several images of tissue stained with Alexa Fluor 488, and I need to determine difference in fluorescence intensity between my control and sample group.
The image files are .TIFF.
Huge thanks in advance!
I would like to image differentiated Caco2 cells grown on a transwell and stained with fluorophore (ie Phalloidin -Alexa 488) without perturping the 3D morphology of the cells.
I have read that you can cut with a scalpel the filter and place it on a glass slide. I gess I should than add antifadding, but then? Should I add a coverslip? Will the coverslip not scratch out the apical surface of the CaCo2 cells?
Do you have a protocol to recommand? any suggestions are welcome
I have tried to fluorescently label graphene oxide sheets once with Fluorescein sodium salt and another time with Fluorescein isothiocyanate (FITC) by mixing 500 uM of Fluorescein (any of them) with 0.5 mg/ml GO overnight then washing with DI water 3X and then re-suspending the GO in DI water. After washing I take a drop of the suspension and place it on a glass slide, leave it to dry then look under the optical microscope using blue light (of wavelength 450-495nm) to excite it but I can't see it fluorescing while the Fluorescein on its own provide green fluorescence at this wavelength. I will be grateful to obtain any advice regarding how to fluorescently label GO sheets.
I want to create a zero-order Bessel beam. But I am a little bit confused how will I create zero-order?. As far I know the most simple way to create a Bassel beam is with an Axicon lens. So my queries are below-
1. Can I use axicon lens to create zero-order Bessel beam?
2. Does apex angle of the axicon lens play a critical role to create zero-order Bessel beam?
3. Is there any other way to create zero-order Bessel beam?
Please help me. Thank you in advance.
I am interested in understanding how the surface area changes when adherent cells are brought into suspensions. I am looking for data supported responses.
I need expert opinions on this matter as my plant samples are detached during washing step of FISH. I used agarose as well as transparent nail polish for this purpose and to me agarose didn't work at all but nail polish works slightly. It provides a base to sliced plant tissue, when I put my hybridization slide into the washing buffer (50 ml falcon) and then in water bath; after heating up to 48 degree Celsius, the nail polish is no more stick and my plant samples swim in freely in the media. Though I doubt that it may not affect the hybridization but still I am not sure. Can you please share your practical experiences?
Thanks in advance
Chromosome counting using fluorescent microscope....
It doesn't matter the color of the fluorophore or the localization (nuclear, cytoplasm). I just need a good one, because the ones I tested so far didn't give great result. Any suggestions from people with experience in that. Thank you very much for helping.
I am trying to isolate a monoclonal cell population by limiting dilution in 96-well plate. I want to isolate transfected cells which they express GFP. Are the cells reduce fluorescent properties by cell division or it is permanent? Should I mark my green cells in first two days prior to fluorescent disappear?