Confocal Microscopy - Science method

May consider using Permai fluorescence dye.

May consider using Permai fluorescence dye.

May consider using Permai fluorescence dye.

Hello, the simple answer is to determine how many cells you need in order to get an assay window of greater than or equal to 2 with the appropriate positive control.

You can also test various numbers of cells for negative and positive control for each end point then use z prime factor to see what gives you the best performance. Zhang et al, 1999, J Biomolecular Screening, 4:67-73

Hello Haiyao Ding

Calcein AM is a non-fluorescent cell-permeable derivate of Calcein that is widely used in cell viability measurement. The carboxylic acid groups on Calcein are modified with AM (acetomethoxy) groups, which endows Calcein AM with high hydrophobicity, facilitating its penetration through cell membrane. Once inside the cell, AM groups are hydrolyzed by intracellular esterases. The fluorescent Calcein molecule is restored, which is trapped in the cell and emits strong green fluorescence.

Since dead cells lack esterase activity, only live cells are labeled and detected. The fluorescence intensity will be proportional to esterase activity. Calcein-AM has been proved to be both specific and sensitive for detection and tracking of apoptosis in living cells. The preservation of membrane integrity is one of the most significant features of apoptosis with respect to necrosis. In the presence of membrane defects, Calcein leaks out of the cell and the signal also vanishes in the presence of residual esterase activity.

On the other hand, Propidium iodide (PI) which is a red-fluorescent nuclear stain is not permeant to live cells or cells which are dead but still have an intact membrane (such as the primary apoptotic cells). In late apoptotic and necrotic cells, the integrity of the plasma and nuclear membranes decreases, allowing PI to pass through the membranes, intercalate into nucleic acids, and display red fluorescence.

Calcein generated from esterase in viable cells emits a strong green fluorescence with an excitation and emission maximum at 494nm and 517nm, respectively, while PI once bound to DNA has a maximum emission wavelength at 617nm when excited at 535nm.

There is something that must have gone wrong with your reagent or your process. You may have cells that are either alive or dead, but not both. Cells which are dead but still have an intact membrane (like the primary apoptotic cells), PI is not permeant to these cells.

You may repeat the experiment. Initially, observe the cells in bright field. Then observe the cells in the green fluorescence channel. The live cells will be stained by green, fluorescent Calcein. Follow it by observing the cells in the red fluorescence channel. The dead cells will be stained by the red fluorescent, PI. Then finally you merge the image of green and red channels.

Good Luck!

Are you sure that this is an artefact? If the antibody epitope is on the intracellular domain it may be genuine signal.

This paper talks about its translocation:

Hey Buse, you could also use dPCR to detect CDs and other markers on the EV surface. I guess that is easier compared to staining, and you will not need too much antibody for it.

You can find the application note for it here: https://www.actome.de/downloads/Actome_EV_Quantification_App_Note.pdf

BLG-nanogels concentrate best at the interfacial boundary of dispersed systems within a pH range of 2.5 to 3.5 because at these pH levels, the BLG proteins carry a positive charge. This positive charge enhances their ability to stick to the negatively charged parts of the interface, helping them effectively prevent the mixture from separating. Outside of this pH range, the charge balance changes, reducing their effectiveness and causing the emulsion to separate more quickly. Essentially, in that ideal pH window, the conditions are just right for BLG to anchor securely at the boundary and stabilize the emulsion.

When staining mitochondria in dead (non-viable) multicellular organisms, traditional dyes like MitoTracker, which depends on membrane potential, are not effective. Here are suitable options for labeling mitochondrial structures in dead cells: MitoTracker Green FM: Stains mitochondria in both live and dead cells, independent of membrane potential. and MitoView Green: Labels mitochondria in both live and fixed cells by binding to mitochondrial proteins.

Epi-fluorescence microscopy (including confocal) will work the same. Just like with cells on glass surface.

Of course the fact that your sample is not transparent will make it harder to find the cells and focus, and the strong reflection from the metal surface might cause issues if the fluorescence filters are of poor quality.

Obviously, you still want to use a coverslip, just like with samples on a glass slide!

May consider using Permai fluorescence dye.

Thank you Dr. Woo.

Hi, I have more to add on this question, how can we prepare such transwell inserts for histology, the ones I am using have an area for 0.3 cm2 ....

May consider using Permai fluorescence dye.

In general, if antibodies work in tissue (confocal), they should always work in cell suspension (flow cytometry), but not vice versa.

Reasons are below:

1. Epitope availability. Some epitopes on the protein target might be blocked in tissue that cannot be accessed by antibodies using the same epitope. Then you may need to go through the "finding the right clone" process to make it work in tissue.

2. Fluorophore limitation. Many scopes are limited by the choices of laser they have, most don't have UV laser in 350nm, therefore you can't use BUV dye conjugated antibodies. BV dyes can be excited at 405nm but they are large polymer dye that may not be staining complex cell processes and tissue structures as well as small dyes such as Alexa Fluor dyes. Many flow antibodies are designed in these colors and not available in other dyes such as BD antibodies, while many tissue antibodies are limited in colors.

Hello, here's a link to a video that shows how to count co-stained cells with ImageJ https://www.youtube.com/watch?v=Z9-Bb68t6ns If you find it helpful, please subscribe to my channel. Subscribers like you encourage me to grow my YouTube channel. Thanks

Dear Albrakati,

I would say any image you acquire should store a value in each pixel (eg: from 0-255 for 8-bit images).

In FIJI, you can open these images and select an ROI (region of interest - a rectangle, polygon, or circle where you want to measure).

Then you press M (measure - Menu > Analyze > Measure). This should pop up a Results table in another window.

PS: you can select your desired measurement parameters in Menu > Analyze > Set Measurements...

Cheers from Portugal,

Vítor Yang

The green signal outside the mCherry-expressing cells is most probably from flavins. Its intensity looks a bit higher than I would expect to see considering similar power densities for all fluorescence channels. But we do not know exactly what laser sources were used. If the green laser was an Andor HLE (2 W), then its power density at "50 ms/20%" would be more than 10-fold higher than that from an ILE 637 nm laser.

Concerning the green signal co-localized with the red one from mCherry, I am pretty unsure that it could be attributed to flavins. I'd rather suppose that you observed fluorescence of the mCherry proteins with incomplete chromophore maturation. It is a widespread phenomenon among RFPs (see, for instance, here 10.1007/s43630-021-00060-8) to show some minor green and/or blue populations. Moreover, fixation procedures can likely chemically modify the red chromophore and thus lead to appearance of green fluorescence.

Hi Masoud,

I would try with some autofluorescence quencher like True Black or Sudan Black. Applied before doing the staining should reduce the background without affecting the intensity of the target areas.

In your experiment focused on studying ubiquitin transfer using confocal microscopy, it's crucial to choose a fluorescent tag that not only provides reliable visualization but also minimally interferes with the protein's activity. Considering your concern about the size of the GFP tag potentially affecting protein function, here are some alternative small-sized fluorescent tags that you can consider for studying in vivo protein activity:

1. mCherry Tag: The mCherry fluorescent protein is relatively small and has been successfully used in various live-cell imaging studies. Its compact size may help reduce interference with protein activity compared to larger tags.

2. HaloTag: HaloTag is a small protein tag that forms a covalent bond with ligands, allowing for specific and reversible labeling. This tag has been utilized for in vivo protein imaging and tracking.

3. SNAP-Tag: Similar to HaloTag, the SNAP-tag system enables specific and covalent labeling of proteins. Its small size makes it a suitable candidate for minimizing interference with protein function.

4.TurboGFP: TurboGFP is a brighter and more stable variant of GFP. While still larger than some of the other small tags, it offers enhanced fluorescence properties and has been successfully used for protein labeling.

5. mEOS: mEOS is a photoconvertible fluorescent protein that changes color from green to red upon exposure to UV light. This property can help distinguish between newly synthesized and pre-existing protein pools.

6. sfGFP: Superfolder GFP (sfGFP) is a compact GFP variant that folds efficiently even when fused to other proteins. It's small size and robust folding characteristics make it a potentially suitable tag for your in vivo studies.

7. Dendra2: Dendra2 is another photoconvertible fluorescent protein that shifts from green to red fluorescence upon exposure to UV or blue light. This can be useful for tracking protein dynamics over time.

8. Blue Fluorescent Proteins (BFPs): BFPs, such as TagBFP, are smaller than GFP and provide a distinct color for multicolor imaging experiments.

Before finalizing your choice of fluorescent tag, it's recommended to perform pilot experiments to confirm that the tag does not significantly affect the ubiquitin protein's activity and behavior. This can include assessing protein localization, interactions, and function in the presence of the tag.

Alternatively, if you're open to alternative methods, you could consider using techniques like proximity ligation assays (PLA) or Förster resonance energy transfer (FRET) to study protein-protein interactions and ubiquitin transfer without the need for direct fluorescent labeling. These methods can provide valuable insights into protein dynamics while minimizing the impact on protein activity.

Dear Manisha,

In my opinion it is not a good idea to check xo-localization with a regular fluorescence microscope.

The staining of your proteins can come from two different focal planes, which means, proteins are not co-localized but look as if they were.

For membrane proteins it should be even more complicated , since you will have fluorescence from the membrane marker all over the cell.

Therefore, use a confocal microscope.

Good luck,

Sebastian

Hello,

I usually incubate my immunofluorescence specimens in a 4% paraformaldehyde solution for 15 minutes and I have never faced any problems following this prodedure (I use about 200 to 250 μl for each slide). I store fixed cells at 4 degrees Celsius in PBS solution containing sodium azide until staining. I've tried keeping them for about 8 weeks and it worked just fine. Just make sure they stay hydrated and sealed to avoid contamination.

Hope this helps,

Best regards,

George

Hello Daniel,

a quick work of advice, squeezing light from a big incoherent source into a small fiber is not only difficult, it is actually physically impossible. This is due to the rule of étendue, which is the optical manifestation of the second principle of thermodynamics.

While if you look up the rule of ètendue you get complex definitions talking about solid angles and apertures, the main, most important consequence of the conservation of étendue is that, given an incoherent source such as an LED or the output of a liquid wave guide, the light intensity per unit of surface can only decrease when passing through a passive optical system. This means that, with your 5mm diameter wave guide, and your 0.4mm diameter multimode fiber, you can at best couple in the fiber a fraction of the light equal to the ratio of the areas, so (0.4^2)/(5^2)=0.6%. You will not be able to couple in anything close to 20%, no matter how complex your optical system is.

Sorry for giving you the bad news, i would suggest looking into multimode lasers for your application, or finding a way to illuminate the disk in your spinning disk without passing through the FC port

Dear Leonardo, I'm also interested in using your tracker! Looking forward to receive news from you

Dear friend Carlos Alejandro Bello Rodríguez

I think you need to connect with your library or google scholar or approaching the authors will certainly help you

Here's a simple protocol you can try:

Hi Oishika Chatterjee

Probably You should try the Prolong Antifade reagent from Thermo (Link Attached). in my experience I have observed that it gives better signal than the Vectashield, Fisher and Sigma's.

I have had some successs treating it like a protein gel asset. I use a small spatula get in between the seal and then slowly twist to crack the well from the slide. You may need to do that in a few spots before fully detaching the wells. This has worked most of the time, but I have had a few break on me if I use too much force. I hope that helps.

Hello,

When performing the APEX2-Gold method on animals, there are a few things to consider:

Overall, the APEX2-Gold method is a powerful tool for detecting and analyzing APEX2-tagged proteins in electron microscopy. With proper technique and high-quality reagents, you should be able to detect a strong signal in your samples.

Regards/

Hi, i think you can use a concentration of 2ug/mL.

I would try confirming the expression by western blot using both myc antibody and antibody for the protein of your interest. The existence of mRNA does not mean that protein was expressed.

Myc tags are not ideal for microscopy because endogenous c-Myc could also be detected and could interfere with the analysis but it usually doesn't.

Kayli Rose Jablonski I suggest to read these articles:

1. https://labanimres.biomedcentral.com/counter/pdf/10.1186/s42826-020-00044-2.pdf

2. https://www.nature.com/articles/s41467-020-19827-1.pdf

3.

4.

5.

6. https://protocolsmethods.springernature.com/posts/combined-whole-mount-fluorescent-in-situ-hybridization-and-antibody-staining-in-zebrafish-embryos-and-larvae

7.

Hello Deven Gopinath

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.

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.

Thanks Both Jiasong and Ayse for your answers!

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.

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

Thank you Alibek, I will do it and I hope it will work :)

Yes. Here is the protocol that we used for detection of proteins on the surface of Lactococcus lactis bacterial cells.

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.

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 ^^

Dear Omar,

do you got the answer for this question? I've meet the same situation and tried to find the explanation.

Thanks.

Hi Diego Suarez have you been using this probe at the end?

I think that you can use the Golgi staining to measure the density

Alexa Fluor 647-coupled antibody, Alexa Fluor 488, Alexa Fluor 500, and Alexa Fluor 514 may might be helpful.

Thankyou

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!

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

Mohamed Khashan

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!

Dear Teena Bhakuni,

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!

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.

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/

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.

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:

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.

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?

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

Dear Chittaranjan Behera , 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.

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

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

Thank you so much.

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:

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

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?

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.

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.

Brian Sanz

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!

You may also contact Thermo Fisher Scientific: https://www.thermofisher.com/sa/en/home/life-science/protein-biology/protein-expression/mammalian-protein-expression/fluorescent-protein-vectors.html

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

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.

There is also a nice online tool to calculate the Nyquist rate:

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.