Science method

Confocal Microscopy - Science method

Confocal microscopy is an optical imaging technique used to increase optical resolution and contrast of a micrograph by using point illumination and a spatial pinhole to eliminate out-of-focus light in specimens that are thicker than the focal plane. It enables the reconstruction of three-dimensional structures from the obtained images
Questions related to Confocal Microscopy
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In my imaging of mouse brains, there are fluorescent dots that appear in the image from confocal microscopy which are not any of the target cells in my experiment. I was wondering if using a secondary antibody from a donkey and a normal donkey serum in the blocking could potentially cause this?
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You should avoid using blocking serum from the same host species as your primary antibody that is detected by a secondary antibody against the same species (immunoglobulins in the blocking serum will compete for the secondary binding).
I agree with Yulia Panina. Washing is an important step. Washing after each application of antibody or other fluorescent probe eliminates antibodies with lower binding affinity present in the sample and thus reduces non-specific signal, or cross-reactivity. Washing for a few minutes in PBS with at least two buffer exchanges will help to eliminate unbound and loosely bound antibody from the sample.
If you are using donkey secondary, then you should use donkey serum for blocking which you have already been doing. Instead of donkey serum you may also try using using BSA (3-5%) for blocking.
Best.
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Dear Fellow Researchers,
We are analyzing adaptive immune response in bladder cancer tissues using immunofluorescene stain and confocal microscopy.
Setting pinhole at AU=1 (perfectly confocal) results is very low signal. Increasing it allows to collect more signal but may yield some over-emission.
What pinhole is acceptable for tissue sections?
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Increasing the size of pinhole will definitely increase the background. I would suggest to keep the pinhole at 1 or may be increasing the pinhole size to 1.5, but in addition, I would definitely suggest to take multiple z-stacks to get a composite image with bigger signal and low background. Hope this helps. Good luck.
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Hi all,
I am growing some large (1mm) organoids in hydrogel. I am planning to stain the organoids for 3 markers + nuclei and image them by confocal covering all 4 channels of UV, green, red and far red. I would like to use the UV channel for staining the nuclei. I am using a clearing agent (RapiClear) clearing the tissue before the imaging in order to be able to see deep inside the structures. The problem is that when I use DAPI, after clearing, the DAPI seems to fade and is not detectable anymore by confocal. Does anybody have experience with SYTOX blue from Thermofisher for staining the nuclei for confocal? Is it stable after clearing the tissue?
Thanks a lot in advance!
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Thanks Both Jiasong and Ayse for your answers!
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Hi everyone.
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.
Stefano Falone
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Hello everyone!
Looking for some useful tips for fluorscence/confocal microscopy of plant root tissue.
I am mostly working with the root tissue of Arabidopsis thaliana where I take confocal/fluorscence images of plant root samples expressing proteins of interest with GFP/mcherry tags also often with Propidium Iodide/DAPI stains. Usually, I don't fix the tissue and take images as such and here are a few problems I face most of the time and I will be glad if you can advise me some suggestions to fix or minimize the following issues-
1.The amount of protein expression in various layers of root tissue is variable ranging from low to very high expression so in a Z-stack often I get highly staurated layers or if I adjust laser power/analog gain to minimize that oversaturation I end up losing signal where protein is less expressing. What should I do in such case?
2. how do you prepare live tissue for imaging, like if you use any specific buffer that does not change the physiological state of the plant tissue but helps in maintianing the GFP fluorescence for a longer duration?
3. Also, no matter how carefully I prepare the slide I never get the root tissue lying uniformly over the slide which means I get regions with good focus and in perfect plane wherease at some regions it gets blurry, out of the focus, how to avoid it?
4. I too have problem with DAPI staining of the root tissue as in most cases barely I see any nuclei but most often I end up with images showing DAPI at the cell borders just like Propidium Iodide.
Thanks a lot in advance for all the incoming beautiful suggestions.
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I work mainly with mammalian cells and have no clue about the plant cells but I feel this could help you with your DAPI problem. DAPI is not great at staining a live nucleus. DAPI most often cannot enter cell membrane/wall and staining requires fixation and permeabilization. I suggest you try Hoechst 33342 which has similar excitation and emission wavelength so you don't have to change your protocol.
Please note that this works for mammalian cells and may or may not work for plant cells but something to think about.
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Dear all,
I am using confocal microscopy to visualise membrane proteins of oleosomes (in any case proteins surrounding the oleosome) extracted from rapeseed.
The oleosomes have been stained with Nile red to detect the oil, and Fast Green FCF (0.013%) to detect the proteins.
The core oil is easily visualised but I am struggling to clearly visualise the proteins which would surround the oil droplet.
Do you have any advice on how to improve the visualisation? It may depend on the microscope settings I guess.
Thank you very much to anybody who could help me
Filippo
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Thank you Yulia Panina, I did more tests and the results are better (no tail and green ring around the red droplet), thus it seems to be matter of sample preparation.
Probably if pH is too high, the FCF dye loses its property.
Many thanks for your interest.
Best wishes,
Filippo
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Hello everybody,
I am doing immunocytochemistry staining for splenic B cells from WT and Knockout mice for one protein. By FACS analysis, it's clear that the signal for the knockout protein is nearly absent in Knockout cells, but when I do confocal microscopy experiment for the same cells, the signal seems to be the same in knockout as in WT. Does anyone have an explanation or can recommend what to do?
P.S: I use the same primary antibody for IF and ICC and No background for the secondary antibody in the control.
Thank you!
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Thank you Alibek, I will do it and I hope it will work :)
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Hi Everyone!
I would like to test by immunostaining that a specific membrane protein is present on my bacteria's outer membrane.
I cannot use FACS for this purpose, so I thought of confocal microscopy as an alternative. I would just fix the cells to keep their membrane intact.
I have not found yet any methods for this. Could you recommend one?
If you have any insight please share it with me!
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Yes. Here is the protocol that we used for detection of proteins on the surface of Lactococcus lactis bacterial cells.
Immunostaining of bacteria for fluorescence microscopy
20 μl of cell culture in stationary phase (OD600=3) was added to 500 μl of Tris-buffered saline (TBS; 50 mM Tris-HCl, 150 mM NaCl, pH 7.5) and centrifuged for 5 min at 5,000 × g and 4°C. The pellet was resuspended in 500 μl of TBS, and 1 μl of FITC-conjugated human IgG antibody (Jackson ImmunoResearch, West Grove, PA) was added. After 2 h of incubation at RT with constant shaking at 550 rpm, cells were washed three times with 200 μl 0.1% TBST and resuspended in 300 μl TBS. Stained cells were fixed to a microscope slide by use of a StatSpine Cytofuge 2 centrifuge (Iris Sample Processing) for 10 min at maximum speed. An LSM 710 confocal microscope (Carl Zeiss, Oberkochen, Germany) was used for observation of prepared specimens. Alexa Fluor 488 was excited with an argon laser (488 nm), and the emission was filtered with a narrow-band 505- to 530-nm filter. All images were taken with the same settings and were analyzed using Carl Zeiss ZEN Lite 2012 software.
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For my project, I need to find colocalization of my retrograde tracer (tiny red retrobeads) with dopamine neurons in (TH immunolabeling, alexa fluor 488).
My problem is that retrobeads are so tiny and I only have few dots of them inside cell. Also finding very sparse red beads in red autofluorescence background in VTA is really difficult under confocal microscope. Does anyone know how can I solve the autofluorescence issue or potentiate my tiny red dots to be distinguishable in red background?
I fix my tissue in PFA 4% and thickness of my slices is 50um.
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Hello,
once you do image treatment, you can define a background treshold on the read signal you find as background. You can also do a Z-stack to enhance your bead signal and also have bead signal in another stack that you might be missing. You can merge your stacks to have the maximum signal. If not, you can also do FACS and seek for double tagged populations ^^
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I'm using JC-1 to measure the mitochondrial membrane potential by laser confocal microscopy. I used 100uM of FCCP as a positive control. Surprisingly, it caused lower green to red ratio and was significantly lower than the untreated cells. My cells are cryopreserved mice oocyte. On the other hand, FCCP did work and caused an elevation in calcium using Fluo 3 AM and caused plasmalemmal membranes depolarization.
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Dear Omar,
do you got the answer for this question? I've meet the same situation and tried to find the explanation.
Thanks.
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Hello.
I need to mesuare the change in quantity and size of lysosomes in different experimental condition and I have this fluorescent dye that stains acidic compartments (LysoTracker Green DND-26) in live cells but It´s used for confocal microscopy So I need to know if someone has ever used in flow citometry with good results and if that´s the case: Could someone give me a protocol ?
Thank you
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Hi Diego Suarez have you been using this probe at the end?
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I was thinking about VGlut2 (presynaptic) and PSD95 (post-synaptic) for the excitatory synapses, and VGAT (presynaptic) and Gephrin (post-synaptic) for the inhibitory ones.
Should I define as a synapse only the areas of colocalization between pre and post synaptic marker? Or should I consider also the isolate VGlut2/VGAT and PSD95/Gephrin ?
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I think that you can use the Golgi staining to measure the density
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Dyes for visualizing bacteria
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Alexa Fluor 647-coupled antibody, Alexa Fluor 488, Alexa Fluor 500, and Alexa Fluor 514 may might be helpful.
Thankyou
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I am trying to analyse the state of the gut epithelium by analysing by confocal microscopy the state of some adhesion proteins (to note, Claudin and occluding); however, these proteins are not exclusive of epithelial cells, which is why I would like to add a marker specific for this subset. Could you please recommend me any marker/combination of markers that makes easier the identification of epithelial cells? Thank you in advance
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Hi Maria,
I agree with other people that EPCAM and E-cadherin are good markers. The staining will be found at the lateral membranes of cells (E-cadherin has strong signal at adherens junctions and weak signal at lateral membrane). ZO-1 is also a good marker for tight junctions although it also stains blood vessels and mesothelium (You can distinguish them by morphology).
When you optimize your protocol, I would reccomend to try different fixation method. Most antibodies of tight junction proteins give great staining with ethanol (or methanol) fixation. Acetone treatment following ethanol fixation sometimes improves S/N ratio.
Good luck!
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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.
Code is:
title = getTitle();
run("Split Channels");
one = "C1-" + title;
two = "C2-" + title;
three = "C3-" + title;
four = "C4-" + title;
run("Merge Channels...", "c1=["+one+"] c2=["+two+"] c3=["+three+"] c4=["+four+"] create");
run("RGB Color");
selectWindow(title)
close()
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I Just checked your macro and it looks okay to me. Maybe you should try and include in the last step
run("Flatten");
I sometimes experience errors with merged images in TIFF format and this seems to help
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We would like to use an enzyme tagged with eGFP and eGFP-tagged alpha synuclein at the same time to observe them under confocal microscopy. However, this will give wrong readings, so we would like to replace eGFP with mCherry. There is a research used the same way but without a detailed explanation.
I will add the link of this research below:
Dinter E, Saridaki T, Nippold M, Plum S, Diederichs L, Komnig D, Fensky L, May C, Marcus K, Voigt A, Schulz JB, Falkenburger BH. Rab7 induces clearance of α-synuclein aggregates. J Neurochem. 2016 Sep;138(5):758-74. doi: 10.1111/jnc.13712. Epub 2016 Aug 4. PMID: 27333324.
thanks in advance.
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As has been mentioned, a simple sub-cloning should do the trick. The caveat being that you have some DNA carrying the coding sequence of mCherry.
Cloning options:
1. Restriction enzyme-based cloning
2. Restriction-free cloning - rf-cloning.org (this works great and is often cheaper, especially if you don't want to order restriction enzymes)
3. Gibson cloning
Happy cloning!
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Hi,
I want to quantify the cell area of bEnd.3 cells (mouse brain endothelial cells) using confocal microscopy by ImageJ. Most of the available protocols need to ignore neighbouring cells where cell-cell boundaries are in touch with each other. However, endothelial cells show continuous morphology. I will be really grateful if someone can help me with the protocol for cell area calculations using ImageJ.
I also want to quantify the distribution of tight junction proteins in cytoplasm vs. plasma membrane of these cells. If there is any way by which this quantification is possible using ImageJ then please help me.
Any help is appreciated.
Thank you in advance.
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If you visually cannot distinguish single&full cells, even with lower magnification, then I am afraid that you need additional staining that evenly stains the membrane to quantify cell surfaces (as I have mentioned above) so the boundaries between cells are clear. Also, nuclei staining would also help to distinguish single cells. PS: General rule: one can only quantify things that are visible.
Regarding the delineation of membrane and cytoplasm compartments - same rule. You can only measure/delineate if you see it, so I am afraid that lower magnification than what you showed above, would not allow that. Also, as already mentioned you have no marker for the membrane, so the delineation will not be very accurate. You would need additional staining of the membrane on top of your protein of interest to be accurate.
Furthermore, is there a way to seed the cells at lower density? This could greatly help your analysis as maybe then you can easier distinguish single cells.
Good luck!
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Can we use deionized water instead of mounting medium for confocal microscopy to preventing photobleaching? Since all the parts of the tissue does not mix well with thick mounting medium and therefore micro-air bubble develops.
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Mounting media usually contain anti-fading agents, so they help prevent photobleaching. If it is too thick, you could dilute it with some PBS. Of course you could also use plain PBS, if you are in a hurry and you fo not care about keeping the sample in good condition for later. Ideally you should put the medium on the one side, place the coverlip at a 45 degree angle, then slowly lower it so that the medium goes everywhere on the tissue with as few bubbles as possible.
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We are going to upgrade our NIKON A1 confocal system with the incubator for long lasting live cell imaging - the incubator should contain atmosfere humidifier, CO2, temperature control etc. Please share your opinions on what system do you consider best and why? What to avoid ? What may be beneficial? The microscope will be used not only for live cell imaging so the system should be reconfigurable . We are looking for something from the top shelf but easy to maintain sterile environment and user friendly.
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I would say that the question doesn't stand Okolab vs. Tokai Hit. There are many solutions from other companies as well.
Recently I wrote a blog post for FocalPlane about temperature control in light microscopy. I think it could be useful for some people reading this thread.
disclosure: I work for a company that produces a temperature control device. It's called VAHEAT and it could be the device for you, if you do a lot of high resolution microscopy, need high temperature precision, need temperature cycles, not just a constant temperature, use microfluidics and more. interherence.com/vaheat/
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I am doing a mammalian oocytes maturation, followed by nuclear meiotic evaluation using confocal microscopy, I mount the oocyte over 90% glycerol in a double sided adhesive tape, and I am curious because I don't want to smash the oocytes under the cover slip. is there any recommendation for a good commercial wax cushion to be used in this case. (oocyte diameter is 100 micron).
Thank you,
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Yes. It is matter of concern and although alternative methods are available in order to prevent oocyte damage during handling in vitro. The alternative methods have some or other type of negative impact on the oocyte. The development of an improved method/materials that can replace glycerol and prevent damage under experimental condition is required. To the best of my knowledge no such appropriate wax cushion is in the market which can be better in handling oocytes in vitro or for the assessment of maturation process in mammalian oocytes for confocal microscope.
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Hi,
I transfected HEK cells with fluorescent ABCG2 transporter (multidrug resistance transporter) , it has a GFP probe attached, and I am performing a live cell confocal microscopy experiment, where I would like to determine the kinetics of Mitoxantrone (chemotherapeutic, far red fluorescence) accumulation in the plasma membrane of the transfected GFP-positive cells. I am doing it by measuring far red fluorescence over time. For the information, mitoxantrone is the substrate of this transporter and ABCG2 is pumping it out of the cell, but when I apply an inhibitor to it, the accumulation rate increases, proving that the inhibitor works. That is how I am testing some of the compounds designed to be potential ABCG2 inhibitors.
Now that I hopefully explained the idea, I can get to the quantification part. I am confused about how many cells/fields of view to use to have a considerable number of sample for quantification. How many transfected cells should I take picture of? 50, 100? Is it for example 100 cells in 3 different experiments, or it is 100 cells per experiment, 3 times ?
As I have to take pictures in certain time point, I can only have one field of interest, where I usually have around 15 transfected cells. How can I collect 100 of them?
Or should I use regions of interest, few of them per one cell, around the membrane and measure that? I am really lost, so please those with similar experience, help!!!
Kind regards,
Marija
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Hi Marija,
People tend to do one of two things when determining sample size. You can either refer to a number of replicates that are „conventionally accepted” in your field for a particular experiment, or alternatively use statistical calculations to determine an ideal sample size based on certain assumptions/preliminary data. In the absence of standard protocols, you could go down this second route. For this you’ll need to have some idea of what an „average” set of results from your experiment look like. The number of replicates and fields of view per replicate will depend on:
  • the variability between your cells/regions within an experiment
  • the variability between independent experiments
  • the effect size (ie how big of a difference) you want to detect.
I would recommend you have a brief read (or watch this YouTube video https://www.youtube.com/watch?v=clbg10IwzYA) about power / sample size calculations so you get the theory, and then try an online calculator such as the one found here: https://www.stat.ubc.ca/~rollin/stats/ssize/n2.html
For the first point, the number of fields of views, the main question is whether your 15 cells you currently image accurately represent the entire well. You would need multiple fields if for example you notice that within a single well there are regional differences. If Mitoxantrone intensity is fairly evenly distributed I would not worry too much. If you are concerned, I was wondering if you could extend your field of view. If you quantify average intensity in the whole cytoplasm, you could probably get away with using a lower mag objective to capture a larger field of view and higher number of cells.
Regarding the variability between experiments, if you have already performed it a few times you should have an idea what the standard deviation is and you could plug that into the sample size calculator.
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Using confocal microscopy imaging technique is it possible to calculate the percent apoptotic index? In some research papers it is mentioned but up to what extent it is reliable and is it necessary to validate using FACS analysis?
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You have to count manually this way, which you may want to do on a regular fluorescent scope first then use confocal to collect your publication quality images. You will quantify this by calculating the percentage of Annexin-V positive and PI-negative cells divided by the total number of cells. Flow is calculating by sorting and is not necessary. Also, something to think about, are you trying to differentiate between apoptosis or secondary necrosis?
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Context:
  • Isolated red blood cells (protocol in .docx is attached)
  • Did confocal microscopy (stained RBCs with CellMask Red (photoiodide))
  • Observed red blood cells (circled in blue) and also some cells with spiky surfaces (circled in green) (image as attached)
Questions:
  • What are those spiky cells?
  • Is it normal for my red blood cell extracts to have them?
  • If not, did something go wrong in my sample preparation, and how do I get rid of them?
Thank you!!
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The simplest explation is that your sample is drying out. As the buffer becomes more hypertonic RBCs undergo crenation. https://www.thoughtco.com/crenation-definition-and-example-609188
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If there are two different Nps prepared for two different drugs encapsulated in a single polymeric system(co-delivery) then can the uptake of both the NPs detected using confocal microscopy? In this case two types of fluorescent dyes needed ??
In another case, for cellular localization expt which dyes except Lysotracker can be used?
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Dear C. B. , the simple and general answer to your first question should be clearly negative, the cellular uptake and cellular location of NPs are different experiments. However, they could be the same experiment if all the uptook NP´s are localized at the same organelle or cell region.
With respect to your second question, if you have two different drugs encapsulated in a polymeric delivery system to facilitate the uptake by the cell, they could be detected by confocal microscopy if these two drugs show distinctive fluorescence signals. It means that these two drugs must show auto-fluorescence in the conditions of excitation used and they must show different "colours" to be distinguished. If one or both drugs do not show fluorescence under the working conditions or they show the same or very close colours, then you must use some dye to label each drug. If your delivery system is intended for reaching some cell´s organelle and just there delivering its cargo, may be it could be possible to just follow the whole system (drug A + drug B + polymeric system) and labelling it with the addition of some fluorescent dye, well inside or attached to the polymeric shell. In this way you could see if the whole system reaches the cell and then the target organelle. To use this methodology you need to be sure that both drugs reach the target and that the polymeric shell keeps its cargo till the destination, in other case you would be following an empty- fluorescent- shell.
Hope this helps. Good luck with your research work.
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how to detect and counted early apoptotic, late apoptotic and necrotic cell population using confocal microscopy?
Is flow cytometry is mandatory to count the population or without using flow cytometry the cell polulations can be counted using confocal microscopy?
For image analysis, confocal microscopy is best of course.
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In apoptotic cells, the membrane phospholipid phosphatidylserine (PS) is translocated from the inner to the outer leaflet of the plasma membrane, thereby exposing PS to the external cellular environment. Annexin V is a Ca2+ dependent phospholipid-binding protein that has a high affinity for PS. Since annexin V staining precedes the loss of membrane integrity which accompanies the later stages of cell death resulting from either apoptotic or necrotic processes, staining with annexin V-FITC is typically used in conjunction with a dye such as PI allows to identify early apoptotic cells (PI negative, annexin V-FITC positive) from dead cells (PI positive, annexin V-FITC positive). you can see this picture for easy understanding- https://www.creative-bioarray.com/support/annexin-v-apoptosis-assay.htm
If you want to distinguish late apoptotic cells from necrotic ones you have to use flow cytometry. You can also check this for reference
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I am interested in studying biofilm formation as a function of multiple variables, and would like to go beyond spectrophotometric/colorimetric/microplate readings and to analyze biofilm structures microscopically as well. Unfortunately, fluorescent microscopy (FM), confocal laser scanning microscopy (CLSM) and scanning electron microscope (SEM), tools that are routinely used by well-funded research groups for this purpose, are currently not at my disposal.
To this end, I am considering to utilize traditional staining (e.g., crystal violet) of these structures followed by simple light microscopy. May I ask for recommendations on objectively/quantitatively analyzing the images that can be generated this way (perhaps through a specific software)? What I have found so far are all for FM, CLSM and SEM images. Thank you!
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Dear Aedrian,
We have faced your problem in the past too - but if you have a good microscope and camera system I think that you can usefully and quantitatively examine differences in biofilm structure using some common dyes for bacterial cells and EPS.
The key questions are whether you have a fluorescent microscope and camera system. If you do, then I'd suggest looking at using live/dead stain mixes, or propidium iodide, and whichever fluorescent DNA and protein dyes you can get. If you only have a light microscope, then you need to be using standard coloured dyes, again for as many cellular components as you can. The second question is whether your camera system can capture single (black and white) or multiple colour images.
No matter what type of microscope and camera system you have, you should be able to take good images of biofilms at a resolution from 10x - 40x and perhaps even 100x (at which point bacterial cells should be visible). You can import these image files into free software such as ImageJ - you can then divide the image into squares and get the total absorbance (e.g. for live/dead stain) and use this in your statistical analysis.
You could look at the variation across the image and correlations between different stains to investigate heterogeneity. You can compare mean values for each image (or measure the absorbance for the entire image) to then compare across images (e.g.different incubation conditions, different strains, etc.).
Regards, Andrew
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I want to perform cellular uptake and localization study of my siRNA-loaded nanoparticles. For performing both the experiments which fluorophore-labeled siRNA, I should order.
Kindly suggest.
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Thank you so much.
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Hi, I am currently working on the colocalisation of 2 proteins and I plan to work it out with confocal microscopy.
In the school I am working we have the leica LasX software with the co-localisation license but it is only available in that one computer connecting with the confocal mic itself. As there are always booking on that confocal mic and the computer is always in-use. I am looking for other softwares for the analysis of co-localisation. Best if I could draw an ROI, shows the pearson's and manders' correlation.
Currently I know ImageJ plug-in JACoP. Any other recommendations? And what's the difference with using the leica LasX co-localization analysis with other softwares such as Image J?
Is there any requirements on the software I use for co-localisation analysis if I am planning to publish my data?
Thanks.
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Hi there,
The main difference of using freeware vs built-in colocalization tools in expensive software is the price :)
Basically, these software incorporate algorithms to express colocalization and pack them into an user-friendly interface, but they're in no way superior to any other (free) software or plugin.
Additionally, you can gain insight into what these software are doing, so you will be able to know which analysis suits better your experiment by using freeware and reading carefully the guides and the manuals (Thresholds, background subtraction, ROIs, and so on...)
I have always used ICA with pretty good results (it operates in a similar manner to JACoP) and you can define ROIs in it. However there are many others. Check the attached documents and links below for practical working examples and explanations. The two attached documents are for ICA, but they provide good working examples and differences for the coefficients
Overall interesting review:
JACop Plugin original paper:
ICA Pugin original paper:
ImageJ forum about differneces in JACoP and ICA:
Why to choose one or another coefficient?
Manders' Original paper:
Guide to ICA:
Another free plugin:
And, for the second question:
"Is there any requirements on the software I use for co-localisation analysis if I am planning to publish my data?"
No, the requirement is not for the software, but for the explanation/justification and interpretation that you give to the analysis and results. Usually, all the Plugins incorporated in ImageJ have been published if that's what you mean
Why you have chosen this analysis? Have you corrected the background? Are you using ROIs? Is this analysis powerful enough? Why you provide one coefficient and not another?
In this case, understanding what you are doing and using proper controls should be enough to validate the analysis tools that you are using for colocalization.
As positive controls, using two secondary Abs against the same 1ary Ab is usually a really good positive control (if you have time and can spend Ab's doing so). You can also perform randomization to see how accurate your coefficients are, but all these things are explained in the attached documents,
Cheers,
J
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Does anyone have any experience/reviews of the EVOS XL Core microscope? 
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Does anyone know how to insert a scale bar on an image using evos xl core? This option is not shown to me on the screen. I’m not sure if it’s not updated or if I have to set this somewhere.
Any ideas?
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(rather than an imaging plate with black wells). Would this work? Or would there be too much light scattering?
Thanks
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If you were to image in an inverted cconfocal microscope, you would need a proper holder that can hold a 96 well plate (depending on what magnification you want to image at). However, you can only image one well at a time. Hence, I would recommend (and you would get much better quality images on) using MatTek dishes that have a glass bottom cover slip.
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I would like to analyze a 1mm in thickness skin sample using confocal microscopy. The sample is too thick to be placed between a slide and coverslip then sealed by nail polish. The idea is to do z-staking and I don't want to cut the sample. Is there is an alternative way to tackle this issue? For example, can I mount the sample using some sort of adhesive tape?
Thanks
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You can add ‘spacers’ as mounts to raise up the coverslip. Use a diamond scribe glass cutter to cut slices of coverslip. Mount these on your slide using nail polish or glycerol in parallel so your coverslip will rest on top of them with your sample in between in mounting media. Then seal round the edges with glycerol gelatine.
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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).
  1. Incubate mouse anti-antigen primary antibody at 4 degree overnight (1:100; used at same concentration as in indirect method staining)
  2. Incubate biotinylated goat anti-mouse secondary at RT for 90 mins (1:150)
  3. 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!
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I was actually going to ask you if you had double checked your filter/Cy3 combination, but since you were already using Cy3 successfully I figured you were good to go. You might throw some of that SA-Cy3 dilution between a coverslip/slide and just make sure you can see it with your 514 set up.
I wish I was being more helpful!
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Hi everyone,
As part of my thesis, I'm looking for gfp-tagged bacteria or other type of fluorescent-tagged bacteria already constructed in order to use confocal microscopy. I've already checked on ATTC but I only found a gfp-tagged pseudomonas. Is there another place (other devices or a laboratory) wich provide that type of bacteria ?
Thank you for the consideration.
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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!
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Brian,
I would not recommend to use MitoSox with fixed tissue!
1. MitoSox is a membrane permeable weak acid. In live cells it distributes into the mitochondria because , as long as the mitos maintain their membrane potential, the mitochondrial matrix is the most alkaline place in the cell. Once MitoSox arrives in the matrix it gives off a proton. Now electrically charged it is no longer membrane permeable which effectively traps it inside the mitos. Thus, for proper localization it depends on the presence of membrane potentials which are missing in fixed tissue.
2. MitoSox becomes fluorescent after reaction with superoxide. The stain is meant to report superoxide as it is produced by the mitochondria, typically by activity of the electron transport chain (normal or pathologic). Fixed mitochondria are not metabolically active any more and therefore should not be able to produce superoxide any longer. Superoxide is diffusible, reactive (ROS - duh!) and therefore has a very short half-life. Thus, the superoxide that has been present at time of fixation will be long gone at the time you stain your fixed tissue slices.
Anything you find with MitoSox in fixed tissue will probably be caused by non-specific effects, such as autoxidation, and will be more confusing than useful.
I recommend to follow Alexandr's hints and concentrate on immunodetection of permanent ROS-damage to the tissue (protein, nucleic acids).
Good luck with your project!
Bernhard
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Hi guys. I'm writing my paper for the first time.
. Representative images of glucose uptake were obtained of wavelengths at Excitation/Emission = 488/516 mm by using confocal microscopy
Is this sentence sound weird?
Or how can i correct this sentence ?
Thanks for nice answering .
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You may use the following sentences:
Confocal microscopy was used to take representative images of glucose uptake. These images were obtained by utilizing excitation/emission wavelengths of 488/516 mm.
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Hi, I'm doing deconvolution to confocal images using Huygens Professional software, and I'm not sure whether I should decide the right signal-noise ratio (SNR) based on the image stack projected by summing the slices (SUM) or by projecting their maximum intensity (MIP). Despite the stack projection obtained by SUM looks better resolved than the original image, it seems to have more background. On the contrary, the projection obtained by MIP looks cleaner in terms of background, but some of the finest structures disappear compared to the original image (look at the attached images). The same is seen by comparing the intensity plots. Then, I wonder whether I should focus in change SNR to obtain better SUM or MIP projection.
Thanks a lot!
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There is also a nice online tool to calculate the Nyquist rate:
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We have a Zeiss LSM 700 attached to an Axio Observer.Z1 microscope equipped with a T-PMT (photomultiplier for transmitted light). When I use the confocal microscope I often want a transmitted light (phase contrast, Ph3) image superimposed on the fluorescence light channel. This gives nice pictures as seen in the first attachment.
However at some point - using the same settings and same protocol - all images from the transmitted light channel has a wave pattern, this is a fault that I do not want in my images. See the second attachment. This problem persists and I have not been able to resolve the problem, do anyone know the cause of this? What I might have to adjust to fix it?
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Hi,
Is this a spinning disk confocal you are using? If so then try increasing the speed of the disk, because the pattern seems like it probably could be forming due to the spinning disk speed being slow. You can get upgradation on the speed.
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Hi,
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
3xPBS, 5min
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)
3xPBS, 5min
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,
Fran
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Hi Fran,
A mild heat-induced antigen retrieval on PFA fixed frozen brain sections has been performed for some IHC assays. At 70-80 C for 30 min or at 40 C overnight in a water bath. Unlike FFPE sections, tissue detachment from the slide might be an issue with HIER on the frozen section Tried several adhesive slides for HIER, Truebond 380 slides found better tissue attachemt or use the floating section method. Good luck
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Hi Everyone,
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?
Thanks.
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Thank you all for your help :) I will read the articles that you have shared.
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Dear all,
I have large, long lipid structures (several µm up to mm) that are present on a carbon coated 400mesh copper grid. My aim is to image these in the TEM.
Using confocal microscopy I can confirm that these structures are bound to the grid and withstand washing and blotting. After each step I check the grid in the confocal and still can see the signal from these structures.
However, a strange effect happens when I negatively stain them( with 2% UFo). In the EM, I can still bright traces (obviously areas without stain) at places where the tubes have been. But the actual tubes are (mostly) not there anymore. There are some small "breaks" in these bright traces where I can see something that has to be a leftover from the lipid structure, but it doesnt seem intact anymore.
I have attached images to better understand what I am talking about.
Has anyone ever seen such an effect and might know how to prevent this from happening?
Thanks everyone for any help/input.
Best
Dario
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Out side my experties
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I am planning long-term studies of tissue remodelling in vitro using light-sheet fluorescence microscopy. I am aware of confocal reflectance microscopy and second harmonic generation microscopy for imaging collagenous extracellular matrix without labelling, but I have seen only "home-made" systems for light-sheet reflectance microscopy. I would like to avoid immuno-labelling collagen, as this would not show collagen formed since the last antibody incubation.
Does anyone have experience in such applications using light-sheet? Would collagen's autofluorescence give a reliable enough signal? I see people are often looking for ways to eliminate this signal, so it seems promising. The quality of imaging doesn't have to be as good as SHG, as that will come later in the project.
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Hi, I was wondering how this project ended up? It seems like a good idea but I wasn't sure if the resolution/signal strength would work out.
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I have cell images from Confocal Microscopy and want to do Nuclear Morphometric Analysis. Can anybody share a detailed protocol to do so?
Thanks in advance.
Swarnali
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If you are good in Python/R there are couple of scripts available. If not, try ImageJ- depends on what depth of morphometry analysis you want to do. You can also use NucleusJ plugin in Image J. Check these paper-
Good Luck !!!
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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.
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Other aspect to think about is the primary ab. What is the epitope recognized by this ab? Some ab are good for western blot but not for IF because of the epitope.
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100- 200 nm fluorescent beads are available from suppliers but only excited at UV (360 nm) or green and red range (i.e., >488 nm). Also. If the objective lens to be evaluated have (NA < 0.8), what is the appropriate bead size to evaluate the PSF from your experience? as 200 nm beads were quite challenging to use for measuring the PSF.
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Hello Ahmed,
I assume you are using a confocal microscope, so you should be able to use any emission wavelength you want, as the resolution should mostly depend on the excitation beam. If that is the case, check the specific excitation spectra of the green-yellow beads you find. While the maximum of absorption will probably be at 488, beads are generally very bright, and have quite wide excitation spectra, so you should be definitely fine using them with a 440 laser, and probably even at 405.
As for the bead size, you should be fine as long as it is smaller than the diffraction limit (wavelength divided two times the NA). The very important part is for them to be very bright, not necessarily big, so in case try to slower your scan speed as much as possible for the measurement.
Also, let me introduce to a very important secret in microscopy: While for published data you should always use the proper calibrated beads from a reliable source, for internal testing you can try to dissolve the ink of Stabilo highlighter pen in water (it has to be Stabilo branded, it never worked for me with pther brands). The ink is slightly hydrophobic, and will form very bright sub-diffraction aggregates. That's a good source of very cheap "microbeads" in different colors.
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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!
Thank you
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Hello, I'm reviving this old post since I'm dealing with a similar situation. I have a recombinant protein with 10xHis tag on the N' terminal and a Myc tag on the C' terminal. The protein is molecular weight is ~42.4 kDa. I would like to use the protein for analyzing antigen-specific memory B cells. I have in mind staining the protein with a fluorophore-conjugated anti-Myc or anti-His, and then using the labeled protein to stain the B cells population, followed by Flow cytometry analysis. Does anybody have an experience with such a procedure aimed at B cells? I am aware of the alternative of using the Molecular probes kits, but the first methods seems cheaper and more straightforward. Thank you!
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Hi,
Has somebody seen this strange aggregation before?
I don't think they are agrobacteria since they are GFP specific (only excitable by 488 nm laser).
Any input is welcome.
Thanks
Alaeddine
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No as I remember I only see expression in the epidermis.
It makes sense that very high overexpression lead to aberrant localization but you still can see the normal localization (nuclear envelope in your case) which is logical.
Any way I will try to reduce the agros concentration it might reduce the expression level.
Thanks
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Certain proteins phase separate into liquid droplets in solution. One can think of these as in vitro membraneless organelles. I would like to find how fluorescent labeled proteins partition inside the droplets using confocal microscopy. I do see intensity change between outside and inside but would like to get some quantitative results from this. I understand I would have to do some calibration using free dye. I am not very familiar with this process. Could someone explain/provide links to how this could be done? Attached is an example of a qualitative intensity profile across a droplet where the labeled protein was concentrated.
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Please check out my paper on multiphase complex coacervate systems on my profile. I detail a Confocal method for measuring fluorescent molecules inside such droplets.
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Whenever I read about confocal microscopy, it is used in conjunction with fluorescence. Wide-field microscopy also has out-of-focus information blurring the in-focus image. It seems that the laserbeam could as well be originating fom behind the sample. Why is it, that confocal microscopy is limited nonetheless to fluorescent applications?
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Confocal microscopy can as well be used to image surfaces or internal structures by recording the scattered or reflected light intensity. Because the light intensity is much higher than with fluorescence, this alows for a much higher scanning speed. Care must be taken that the detectors can cope with these intense light powers. PMTs must probably be protected using a small pinhole setting and minimum illumination beam intensity or a filter.
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I am looking for fixation protocols to non-adherent cells for confocal microscopy analysis. Also, I would like to know for how long I can keep the cells fixed (for example, I want to check if I can fix/prepare many plates during the week and take images from all of them only on Friday).
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José Carlos Quilles Jr You can try the following protocol. Also, if you want to use coating reagents such as Poly-L-lysine, you can coat 0.1 mg/mL of PLL (in autoclaved water) for 5 minutes by shaking. Wash this with autclaved water and dry for 2 hours. You can then add the cells there and fix them. Fixed cells can be imaged after multiple days or week (if the fluorescence is strong enough)
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Hello,
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?
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Hi there,
The answer above is all right and is more of a BatchProcessing. You have also another option which is to assign a shortcut to your Macro so you just have to press a key to perform the function. In the example attached here is "F9". Btw, it will be much faster if you decide a value for each one of the channels for B&C adjustment so you can write it in the macro, this will prevent you for adjusting it in each channel each time. If you want to use different values for different sets of images, just rewrite the macro. The macro here automatically does all what you were doing, but image by image, by pressing F9. It will save the final image in the same folder where your original image was.
To use it:
Download the *.ijm file attached here.
Go to Plugins>Macro>Install...
Then open the image and press F9 and it will be done.
It also creates a white scale bar of 10*1 um at x=2um y=2um of your image. If you want to change those dimensions or the position, just modify the values 10 and 1 in the run("Specify..." line.
One warning, this is assuming that your C1=Red, C2=Green and C3=Blue. If it is not the case you have to modify the run("Merge Channels..." line. c1, c2 and c3 are always red, green, blue. If you have more channels, you will have to add them following the same reasoning.
Hope it helps,
Cheers
macro "adjust BC [F9]"
{
title = getTitle();
filename_clean = File.nameWithoutExtension();
path_to_dir = File.directory;
path_to_save = path_to_dir+filename_clean;
run("Z Project...", "projection=[Max Intensity]");
close(title);
run("Split Channels");
selectWindow("C1-MAX_"+title);
run("Brightness/Contrast...");
waitForUser("Apply optimal Brightness and contrast settings, then press OK");
selectWindow("C2-MAX_"+title);
run("Brightness/Contrast...");
waitForUser("Apply optimal Brightness and contrast settings, then press OK");
selectWindow("C3-MAX_"+title);
run("Brightness/Contrast...");
waitForUser("Apply optimal Brightness and contrast settings, then press OK");
run("Merge Channels...", "c1=C1-MAX_"+title+" c2=C2-MAX_"+title+" c3=C3-MAX_"+title+ "");
run("Specify...", "width=10 height=1 x=2 y=2 scaled");
setForegroundColor(255, 255, 255);
run("Fill");
saveAs("tiff", path_to_save+".tiff")
run("Close All");
}
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I have different titanium surfaces scanned using confocal scanning laser microscopy (CLSM) and need to analyze their roughness (Str, Std, Sdr, and Sds values are needed). thanks in advance for your precious time.
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Thank you all for your quick reply. I started to check them out one by one. Mr. Dirk Luetzenkirchen-Hecht than you but usnforturately Gwyddion does not have hybrid parameters command for the measurements I need (Sdr. Sds, Str, etc). I search and check many time with no results. Digital Surf is also have this lack of parameters unforturately Mr. Márcio Mafra. Thanks again for all.
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I do not have a TRITC filter or other laser set available other than 488nm.
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Dear Akeel M. Kadim, did you just copy-pasted part of my response as your own answer?
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I want to quantify/count neuronal cells, pretty well filled up with the expression signal of a protein. The images have been taken via confocal microscopy at 10x resolution. Which software/application/program is best and accurate to count the cells ? I don't want to count them manually!
Your suggestions can help me to analyse my data efficiently!
Much thanks..
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You should be able to use Fiji (imageJ) for doing these. I use the plugin called QuimP on Fiji ( you can get the QuimP guide online) and it works pretty good for me to do segmentation and analysis (to be precise it gives you area, fluorescence (both cortical and cytoplasmic), circularity, elongation analysis followed by a heat map too if you want to generate one.
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Does anyone know a simply way (i.e. using free image softwares) to quantify chromatin condensation? Cells were labeled with DAPI and analyzed by confocal microscopy.  
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You can calculate the chromatin condensation parameter (CCP) simply by staining the cells with DAPI.
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In addition to clinical examination, dermoscopy and RCM help in diagnosing AHM. Which would be a better option between the two? Why?
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Thanks Khadija Elboukhari for your excellent insight.
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Hello All,
I want to compare the surface roughness among different samples from confocal microscope (TCS-SP8, Leica), for example, to get the Rq or SA value from the surface.
Does anyone know the method to do this, or is there some software to do the calculation?
Thank you in advance.
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Qiran Chris I do not have the machine available, while thanks for your answer.
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I've used the NIKON A1 inverted confocal microscopy a couple of times and I noticed the red stain at 540/565 nm gets bleached within seconds. The objective I use is an oil 60x. Could anybody please let me know how this can be fixed? Massive thanks!
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Dear Tommy,
You can use deblurring and sharpening algorithms to restore the quality images / video. If necessary I can give you the codes.
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We are operating a Bio-Rad MRC-1024 LSCM on an OS2/Warp computer that is failing. I'd like to upgrade to a newer computer and more recent software, but Zeiss has discontinued the LaserSharp software that would run our LSCM, and they have no replacement. Has anyone found an alternative software to run this hardware? Alternatively (and assuming no objection from Zeiss), could we borrow the (now obsolete) software to install?
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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?
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DAPI stains RNA as well, unless you pre-treat the cells with RNAse. Maybe try Hoechst (33458?) because it does not bind to RNA, just to DNA (it binds to the "rungs" of the DNA ladder, which RNA doesn't have).
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Hello, everyone. I am performing scrape loading/dye transfer assay at moment(using lucifer yellow) and get some green fluorescent images for analyzing the Gap Junctional Intercellular Communication. But don't know how to get the dye spread distance and fluorescence area via imageJ software. Is there anyone who could give me some advices? Thanks so much.
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Or you can also use macro for more automated process :)
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Hi guys,
I am trying to culture mouse embryonic stem cells ( JM8A3; C57BL/6N ) on coverslips coated with 0.1% gelatin for confocal microscopy. The cells grow well but once I start washing the media with PBS, the cells detach from the coverslip. Anyone can suggest a way to fix that?
what is the best way to grow mESC for confocal studies?
Thank you in advance
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Dear Yulia , Thank you for your answer , don't u use mounting media after Dapi staining ?
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I am analysing an immunofluorescence dataset taken by a confocal microscope in one z-plane. For each cell and channel, the mean fluorescence intensity in the nucleus and cytoplasm are reported. I was now wondering if the relationship between the mean fluorescence intensity and the concentration is linear.
  1. For instance, if a cell has 10 times as much nuclear mean fluorescence intensity in channel 2 than another cell, can I reliable say that the concentration of the antibody target protein in the nucleus is 10 times as high?
  2. Similarly, if a cell has 10 times more channel 2 mean fluorescence intensity in the nucleus than in the cytoplasm, can I reliably say that the concentration of the antibody target protein in the nucleus is 9 times higher than in the cytoplasm (assuming that ca. 10% of what is labelled as cytoplasm is actually golgi/ER/mitochondria/etc. and that these organelles do not contain the target protein at all)?
Thanks,
Paul
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Usually with a CMV driven vector transiently transfected in to a cell line you will get a huge range of fusion protein expression levels (usually beyond the dynamic range of a single image with most imaging platforms). The expression level will depend on the fusion protein half life, the time after transfection, - or more accurately nuclear access- and the number of copies of the vector introduced into cells. Cells will only start to express protein from the vector after mitosis and the levels will then tend to increase untill the cells are fixed.
Even if your endogeneous protein is cell cycle regulated in abundance then you should still get a good linear relationship at the top end of the expression levels, the regression line will just not go through zero. The non-transfected cells in your population will give you a good idea of the dynamic range of the cell cycle regulated endogenous protein levels. Incidently, after a transient transfection (say after 24hs with fugene6 ) transfected cells usually appear in pairs with similar expression levels. The distance between the pairs of cells can give you a crude estimate of the time since mitosis.
Phospho specific antibodies are more of a challenge - you could try expressing a phospho-mimetic mutant vector to see if that works with your antibody, if it is specific for a particular phosphorylation site?
We once managed to get a really good polyclonal phospho specific antibody to a serine in I kappa B alpha that on western blots was much more sensitive (at least 10x ) than the antibody to I kappa B alpha itself, which just goes to show how varied antibody affinities are.
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Hello!
I'm working on a project in my university. Among other things, I'm required to count the number of astrocytes in the image stacks I've been given, as well as measure the length and thickness of their processes, using Fiji. However I'm having lots of trouble recognizing what might or might not be an astrocyte in each stack. I was told to create a composite image and a max intensity Z project image of GFAP stain (for astrocytes) + DAPI stain (for nuclei) then manually count based on colocalization of the stains, but I find that this leaves me with somewhat confusing images where I'm not sure if what I'm looking at is actually an astrocyte or not.
I've left a Max Intensity Z project image as an example. In this case there are many areas where I'm not sure if it's a full astrocyte or just a part of another one. And when identifying what I could more safely assume is an astrocyte (like the one on the upper right quadrant) I still can't see if there is a stained nucleus that belongs to that astrocyte. I can see there is a nucleus in the same zone, however, it appears to be much bigger than the astrocyte itself (I'd expect the nucleus to be "inside" the GFAP stain, like I've seen in other pictures). Would you for example count that as one astrocyte? And are there any tips to effectively identify all astrocytes in an image?
Thanks beforehand. I'm sorry if this comes off as confusing, but I'm really not sure how to work with this.
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The problem is converting to a Z-projection. This approach will superimpose cell nuclei and cell processes from multiple levels, making a pattern that is confusing at best, and deceptive at worst.
You are far better off manually counting cells by 'scrolling' through your Z-stack, and counting / clicking on nuclei that are definitively enclosed by green cell bodies. FIJI / Image J can then count the sites you picked and even give you the coordinates.
If you have many cells to count, you might be able to segment your image stack into particles based on the target object (blue nuclei surrounded by green) and then count them automatically. But by the time you do all the thresholding and segmenting, you might as well have done it manually-- with fewer errors. Nothing in image analysis works quite so well as the human brain.
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Hello,
I am studying cytokeratin 14 (K14) expression in Hacat human keratinocyte cell line and also von Willebrand factor (vWF) expression in choroid retinal endothelial cells (RF/6A). RF/6A cell line is obtained from rhesus species.
I have used following antibodies for the study:
1. Anti-cytokeratin 14 antibody (Anti-human) raised in mouse - ab7800 (Abcam)
2. Anti-vWF (anti-human with cross reactivity to non-human primates) Monoclonal Antibody (F8/86) raised in mouse- MA5-14029 (Thermofisher)
3. Goat anti-Mouse IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 633- A21052 (Thermofisher)
I have used following protocols for Immunofluorescence (IF) studies.
1. Culture respective cells on coverslip in sterile conditions overnight (until attachment occurs)
2. DPBS washes, fixation (two batches- 4% PFA and Methanol and DPBS washes again
4% PFA fixation: 20-25 mins at room temp
Methanol fixation: -20 deg Celsius for 8 mins
3. 1X DPBS washes (twice), Permeabilization in 0.2% and 1% Triton X 100 (Permeabilization was done only to RF/6A cell line samples fixed in 4% PFA to test for the presence of vWF)
4. Blocking - Conducted for 1h at room temp
a) For hacat, 5% BSA in DPBS was used as blocking agent
b) For RF/6A , two batches comprising of - 1% BSA only and 1.5%BSA+0.5% Triton X 100 were tested for blocking. (This was as per the protocols suggested by different colleagues in the institutes around ours).
5. Primary antibody dilutions:
a) Anti-cytokeratin 14- 1:50, 1:150, 1:250
b) Anti-vWF- 1:50, 1:100, 1:150
Incubation overnight at 4 degree Celsius in a humidified chamber.
6. 1X DPBS washes thrice on a shaker, and incubation with secondary antibody (dilution used 1:500) for 1-2 h (in dark)
7. 1X DPBS washes thrice on a shaker (in dark)
8. Mounting of slides on EverBrite Hardset Mouting Media with DAPI (Biotium) and stored at 4 deg Celsius prior to imaging by laser scanning confocal microscopy.
Lasers used: Excitation - 633 nm
Emission- 647 nm (Tried from 637 nm- 655 nm)
We tried adjusting laser power and intensities. However, we were receiving bright red fluorescence of the background instead of the cells. DAPI fluorescence was observed properly. Even in negative control slides (only secondary antibody), we could only see background fluorescence like the ones attached herewith (negative control images of both 4%PFA and methanol fixation are attached).
Thus, we are not able to study marker expression properly.
I have attached images of:
a) 4%PFA fixed negative control
b) Methanol fixed negative control
c) K14 4%PFA fixed 1:50 dilution (primary Ab)- Hacat cell line
d) K14 Methanol fixed 1:50 dilution (primary Ab)- - Hacat cell line
e) vWF (4% PFA fixative, 1.5%BSA+0.5%TritonX blocking and 1:50 primary Ab dilution) - RF/6A cell line
f) vWF (Methanol fixative, 1%BSA only blocking and 1:50 primary Ab dilution) - RF/6A cell line
Kindly guide me regarding the same.
Thank you.
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I do have some question regarding the protocol you presented:
i) How have you been cleaning the coverslips prior to cell culture? To minimize autofluorescence and minor impurities of the glass for confocal microscopy, you should clean them with strong acid and alcohol prior to sterilization in an autoclave.
ii) You should not consider using methanol as a fixative when investigating intermediate filaments (or other cytoskeleton proteins) for imaging. Please consider using PFA.
iii) When you apply protein crosslinkers, such as PFA as a fixative, you should always treat coverslips with a quencher (glycine or NH4Cl);
iv) Consider using Triton-X in a lower concentration (0.1–0.3%).
I would believe this autofluorescence effect seen during your microscopy is mainly due to the immunofluorescence protocol itself, and not in image acquisition... This can be seen when comparing to your control frames.
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Mechanical contact stylus techniques are the traditional method of measuring the internal diameter (ID) of shafts. Bore gauge is a good example for this method. Now I am trying to use non-contact method and after researching about different non-contact techniques and the available equipment, I found confocal chromatic sensor very accurate (as low as 0.5-2 microns). I learned how to use it to find some GD&T features like run-out but I don't know how to find the bore size (internal diameter). The case study is explained below. Any suggestion is highly appreciated.
Geometry: A cylindrical shaft with internal diameter of 0.926" and length of 40"
Question: To find the size of ID by using confocal chromatic sensor.
Thank you
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Several
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As far as I know, the only difference between a confocal and conventional fluorescent microscope is a pinhole added between the detector and dichronic mirror.And also for the two-photon microscope what we need is just an augmented light source.
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Hi Hasan,
First of all, it depends on what kind of "conventional" fluorescence microscope(s) you have. Specifically, you need a conventional fluorescence microscope of an infinity-corrected design, and this microscope needs to have a light path available for you to add the excitation laser(s) mentioned in above answers.
If that's the fluorescence microscope you have in hand, you can either consult someone who has the experience building a confocal microscope, or get this book 'Handbook of Biological Confocal Microscopy,' which has a few chapters detailing the procedures of building the microscope. If you have plenty of experiences working on free-space optical setups, I would say this should only take you a couple of weeks. If not, this can take months. Also, depends on your hardware programming proficiencies, the hardware synchronization can take you several days to months as well.
Alternatively, if what you really need is optical sectioning equivalent to confocal microscopy, maybe you want to take a look at some simpler designs such as Neil et al.s' work (Optics Letters 22, 24, 1997) or York et al's work (Nature Methods 9, 7, 2012, see its supplementary materials). These designs do not require scanning mirrors or photomultiplier tubes, so they are somewhat easier than conventional confocal microscopes in terms of the construction works and hardware synchronization.
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