Oh, this is a great question - and answer: it really depends on what you want to image. For example, I am looking at trafficking of proteins into the cell wall. The cell wall usually has a lower pH, often around 4.5 -5.5, so I would be looking at fluorophores that have a pKa around that range; e.g. mCherry (pKa 4.5), mRuby3 (4.8), GreenLantern (pKa 5.6), mNeonGreen (pKa 5.7) or Citrine (pKa 5.7) would be great candidates. I also need a fluorophore that folds fast so the fluorophore is functional in the endomembrane system and I can visualize the protein trafficking. mCherry, GreenLantern and mNeonGreen all are predicted to fold within 15 mins, at least at regular cell culture conditions around 37C, while mRuby3 is a slow folder, with more than 2 hours until maturation; so the latter is not as good a choice for this experiment.

The choice of fluorophore for your experiment depends on the microscope you will use, the tissue/cell line/cellular compartment of interest, and the temporal scale of the experiment. Fluorophores are continuously updated by the community, and besides their range of excitation and emission wavelengths, they vary in several important parameters, such as folding time, brightness, lifespan, tendencies to form multimers, and pH sensitivity/pKa, amongst others. These parameters might also vary depending e.g. on temperature or mounting media used for the experiments. For an extended list, please go to: https://www.fpbase.org/.

Fluorophores for single-molecule microscopy can come in several different flavors. Fluorophores can be genetically encoded fluorescent proteins, which are frequently used for live-cell imaging (e.g. EosFP, Dronpa, or PA-mCherry), or various synthetic dyes such as Alexa.

These emitters, when not genetically encoded, can be linked to the molecule of interest using one of several options: immunolabeling with either a molecule-specific primary antibody combined with an emitter-fused secondary antibody or a small antibody fragment such a nanobodies or single-chain antibodies fused to the emitter, or alternatively binding of the emitter to a small protein tag.

The choice of fluorophores/dyes used for super-resolution microscopy depends on the technique utilized. For example, STORM and PALM utilize sequential activation of photoswitchable fluorophores to build a time-resolved, high-resolution image and thus requires the use of photoswitchable fluorophores or dyes, such as Alexa-Fluor dyes with different wavelength specs which can be conjugated to an antibody or other protein for imaging. STED, which selectively deactivates fluorophores, can and commonly use Alexa-Fluors, but can also utilize fluorescent proteins such as GFP, or CLIP-/SNAP-tags for in vivo imaging. For SIM, which combines use of a projected periodic grid to create interference patterns with deconvolution, both synthetic dyes such as Alexa-Fluor and genetically encoded fluorophores such as GFP can be used. 

Some commonly used softwares to track your molecules over time are Imaris, Metamorph and ImageJ/Fiji plugins, such as Trackmate. Here are some protocols describing single molecule tracking in 2D and 3D: https://bio-protocol.org/e4390, https://bio-protocol.org/e3794 and https://pubmed.ncbi.nlm.nih.gov/27713081/

Hello,

Use of a fiducial marker would help you to differentiate between these two movements. A good fiducial marker is basically anything that can be visualized by your system and is completely immobilized on the sample substrate. The immobile object can then be used as a reference object as the image is acquired. The fiducial marker will remain fixed within the sample, but will move if the sample drifts. Images can be normalized/aligned to the fiducial and any remaining movement attributed to the actual movement of the cell. This alignment of the fiducial can be achieved during imaging with a stage that monitors and responds to the fiducial position (as in the webinar) or in post-processing image analysis using image alignment software functions. I hope this answered your question.

Fiji/ImageJ (https://imagej.net/software/fiji/, https://imagej.nih.gov/ij/) are great open-access and community driven resources, with extensive plugin options for more specialized analyses; one example for tracking particles is TrackMate (https://pubmed.ncbi.nlm.nih.gov/27713081/), and you will be able to find many other useful macros/plugins. Another frequently used image analysis software would be Imaris, which comes with a suite of useful applications.

Fiji Image J

HI Qian Luo, that's a great question! I am interested in the localization and dynamics of different proteins in living organisms and am using a spinning disk confocal microscope, that gives you high temporal and spatial resolution (if is quite fast) and can also perform optical sections, so I have used it to determine protein localization patterns in 3D. And depending on the system you're using, you can image a few layers of cells into the organ, which is quite useful.

Renate Weizbauer, thank you very much for your answer! We have spin disk confocal microscope for fast image and it is quiet useful.

Hi William, from my experience, I am using the Glass Bottom Dishes to grow cells for microscope imaging. Different types of glass bottom with different size, such as 24 well-plate or 35 mm glass dishes; coated or non-coated can be purchased from different suppliers. They are really great for cell culture and high resolution imaging.

They are working for both fixed cells and live cell imaging. For the live cell imaging, we have the incubator together with the microscope that controlling the growth conditions, such as temperature, CO2 and sterile. For different aim, I usually culture cells for 24hours or 48hours before imaging. Hope it answers you.

Hello,

that is an excellent question. Here you find a number of publications on the what are good practices in image processing and which ones are not.

https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4114110/

https://rupress.org/jcb/article/166/1/11/34064/What-s-in-a-picture-The-temptation-of-image

Sure. Thank you very much for sharing.

Single-molecule localization microscopy (SMLM) requires a separation of the point spread function (PSF) of multiple individual emitters, which can be achieved by e.g. temporal separation (photo-conversion/switching/activation of neighboring emitters or intermittent binding of the emitter to the molecule of interest) and can enable spatial resolution of ~20-50nm. A variety of microscopes can be employed, including confocal microscopes, TIRF microscopy and light-sheet microscopy, as long as the signal to noise ratio is permissible. (e.g.https://bio-protocol.org/e4390).

HI Chenchen Liu,

this webinar focusses on single-molecule localization microscopy (SMLM), which is one of several super resolution microscopy techniques. It can enable spatial resolution of up to ~20-50nm, which is better than the ~200nm resolution of conventional confocal microscopy. Single-molecule microscopy is based on the ability to separate multiple individual emitters in close proximity. This separation can be accomplished by several mechanisms, including temporal separation (photo-conversion/switching/activation) of neighboring emitters or intermittent binding of the emitter to the molecule of interest. You can use different microscopes with this approach, including confocal microscopes, TIRF microscopy and light-sheet microscopy, as long as the signal to noise ratio is permissible.

HI Divya Bhagat,

there might be several ways of visualizing gene expression - or mRNA levels -, one utilizes an in situ hybridization technique where a single-stranded and fluorescently labelled DNA probe hybridizes with your RNA of interest. That techniques enables quantification of both the amount (quantity) of the RNA and its localization within the cell. Here is a protocol explaining how you can apply this technique:

https://bio-protocol.org/e3070

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https://bio-protocol.org/webinar

HI Rashi,

that is an interesting question. What do you want to look at in microalgae using the microscope? What spatial and temporal resolution do you need? That might determine the kind of microscope you might choose to use.

thats an interesting question. Single-molecule localization microscopy (SMLM) is based on the separation of the point spread function (PSF) of multiple individual emitters. This can be achieved by several mechanisms, e.g temporal separation (photo-conversion/switching/activation) of neighboring emitters or intermittent binding of the emitter to the molecule of interest), leading to a spatial resolution of ~20-50nm, far better than the ~200nm of conventional confocal microscopy. A variety of microscopes can be used with this approach, including confocal microscopes, TIRF microscopy and light-sheet microscopy, as long as the signal to noise ratio is permissible.

Dear Dahou Sara,

thank you for your question. the staining protocol with Fluorescein 5-isothiocyanate (FITC) depends on your approach and organism, there are a number of protocols at bio-protocol.org covering experiments with that stain. Frequently, FITC is used conjugated to an antibody that then recognizes your compound of interest e.g. https://bio-protocol.org/e834

You can also find several protocols covering Netosis, e.g. https://bio-protocol.org/exchange/protocoldetail?id=3927

hi, that's an interesting question/vision.

There might be several ways of visualizing gene expression - or mRNA levels -, often utilizing an hybridization technique where a single-stranded and fluorescently labelled DNA probe hybridizes with your RNA of interest. That techniques enables quantification of both the amount (quantity) of the RNA and its localization within the cell. Here is a protocol explaining how you can apply this technique:

https://bio-protocol.org/e3070

another one, RNA paint, which is based on FISH, can visualize chromosomal regions where genes are being transcribed: https://bio-protocol.org/e1914

These are just some examples to visualize transcription in a cell.