How to add a scale bar after an image acquisition by a confocal microscope (Leica, Nikon or Olympus)?

Two methods are available:

  1. – The image was acquired on one of our confocal microscopes and the file was saved in a proprietary format (Olympus OIF, Leica LEI/LIF, Nikon ND2)

1.1.    Open the file with its viewer (links available on the platform website) and ad the scale bar manually

1.2.    If this method doesn’t work :

1.2.1.  Open the file with ImageJ (be sure that the plugin LOCI Bio-Formats is already installed, link and instructions on the website « links » page)

1.2.2.  Check that a dimension, in microns, is available just below the title of your image window

1.2.3.  Go to Analyse->Tools->Scale Bar

1.2.4. Choose your scale bar settings and click OK

1.2.5. Save your image


2. - The image was acquired with another system (classic or confocal microscope) and you have only the image file (in .TIF, .JPG, …) without scale bar

2.1.    Go back to the same microscope used for your acquisition and acquire a new « blank » image (no need to have a real sample), using EXACTLY the same settings of you old image (magnification, resolution, zoom, …) and be sure to have a scale bar from the system in your “blank image” this time. This image is now called “reference image”

2.2.    Save this reference image using the same format of your old image (.TIF, JPG, …)

2.2.1. If JPG, be sure to use the same compression the old one

2.3.    Open your reference image on ImageJ

2.4.    Go to Analyse->Set Scale and cick « Click to remove scale », then close the window

2.5.    Draw a straight line (you have a button for it on ImageJ  main window) which will represent the whole length of the scale bar (by precaution, you can keep in mind  the value of this length in pixels, showed at the bottom of ImageJ main window)

2.6.    Go to Analyse->Set Scale and you will find this value on « Distance in pixels ». If it’s not showed, you can write it down.

2.7.    Fill the « Known distance » with the real value of your scale bar and “Unit of length” with “microns”

2.8.    Without closing the reference image, open your image (without a scale bar)

2.9.    Go to Analyse->Tools->Scale Bar, choose your settings and click OK

2.10.Save your new image (with a scale bar)


Lenses optical resolution (Optical quality)


We often associate image quality to its magnification. Even if this information is important, there are two other factors which are also important. For optical microscopy, the quality of an image is made by associating magnification, numerical aperture and digital resolution.

Magnification: how many times the image is bigger than looking with your own eyes

Numerical aperture: One of the most important parameters to the optical quality of your image, this value will indicate the resolving power of your lenses, the ability to separate two independent objects. Two lenses with the same magnification but different numerical aperture will give you two different images, mostly affecting the quality of fine structures like membranes, vesicles or spots, but also other structures.

Digital resolution: represent the number of pixels (image unit) on a whole 2D dimension of an image. On optical microscopy, a value of 1024x1024 is many times used by default. It’s important to know that raising the digital resolution more than that will not increase the image quality in most of the cases, and do not forget that raising the digital resolution value will decrease the scan speed increasing the bleach factor.


Supports used in microscopy


Several kinds of supports may be used in microscopy:

Slides – one of the most classic supports for fixed cells, a slide should be always “mounted”, it means that a mounting medium must be used between the slide and a coverslip. This mounting medium has many functions: isolate the sample from the environment, keeps the fluorescence intensity for a longer period (anti-fade), and allows a homogeneous field of view. Several types are available, for example the Vectashield (https://www.vectorlabs.com/catalog.aspx?catID=279), Prolong  (http://products.invitrogen.com/ivgn/product/P7481) or Moviol/Mowiol (http://www.polysciences.com/Core/Display.aspx?pageId=98&categoryId=57&productId=920). Other brands are available and are also adapted to the confocal microscopy, but remember to contact the platform staff before using another brand to be sure that its refraction index is adapted to our lenses.

Petri dish – Classic support for live cells, provided with a plastic or glass bottom, this support allows the sample to be in a liquid environment during time-lapse acquisition. Glass bottom dishes are better to be used because the light path will be less modified bringing more quality to images. Anyway, if the sample must to be in a plastic bottom dish, it’s still possible to use our Nikon A1R which is equipped with “long distance lenses” (20x or 40x). There will be a slight loss of optical quality but allowing the focal plane to be further than with classic lenses. A multi-sample dish with glass bottom is also available, which is compartmented into 4 quadrants allowing doing 4 different conditions at a time.

Multi-well plate – this support is used to acquire a large number of different conditions in parallel. For imaging, the same plastic/glass bottom rule is applied and normally there are several brands available, with a possibility to have from 24 to 384 wells. Some brands optimize a plate to their own equipment (BD has a BD Pathway optimized plate), but a BD plate can also be used with another system, if needed (our Nikon A1R is ready to use multi-well plates and a BD plate can used with this microscope).

Othe special supports:

IBIDI: their  particular supports are made on a coverslip-like bottom with a classic slide or Petri dish dimension to different multiple-conditions adapters in one single unit. Their bottom has a refraction index really close to the glass which allows the researcher to use them with any of our microscopes. Others adaptations allow users to reduce the quantity of reagents used (channel slides) or to use with special applications (like 2D/3D chemotaxis migration assays or scratch tests).


Labtec: their supports are also based in a multiple-conditions assay but the bottom is a real glass slide, so the user needs to unmount the support to be able to use the classic slide attached to the adaptor (as a classic slide with coverslip). Also, they have a ready-to-use “chamber slide” which doesn’t need to be disassembled to observe in microscopy. Just remember that this last one has a different total dimension, so only microscopes with a universal holder can be used (like our Leica SP5 ou Nikon A1R).

Milipore (Millicel EZ slide) : also based on a glass slide, this is an alternative to labtec with different chamber configurations.




HCS – What’s a High-Content Screening and which are the advantages comparing to classic microscopy


This system will allow the researcher to acquire multiple sample in parallel (e.g. in a 96/384 multi-well plate), on a stand-alone system of acquisition, image treatment/analysis and data analysis. The phototoxicity using this system is well reduced due to a double lamp spinning disk confocal microscope. The system is also equipped to time lapse experiences (T°C and CO2 control) as well as an automated fluidics handling control. Some applications of this system are regrouped in the next figure:


Additional information is available at : http://www.bdbiosciences.com/documents/Pathway_brochure.pdf



What is my fluorochrome spectra?


It’s crucial to know the excitation and emission spectra of all the fluorochrome you use for your experiments, to be able to configure the optical path as it should be and acquire very good images. All confocal microscopes at the platform are equipped with spectral detectors. This kind of detector allows the user to configure freely where the detection starts and where it ends to have the best of emission intensity from their sample. If the sample is added of more than one fluorochrome, it’s better to use the most separated possible spectra to avoid overlapping, so the user doesn’t need to use corrections methods (sequential mode or unmixing).

To help users with some classic fluorochromes, a spectra viewer is available at





F techniques  (FRAP, FRET, …)


These are techniques developed to analyze dynamic cellular events.

The FRAP (fluorescence recovery after photobleaching) allow the researcher to determine if a structure is mobile inside a compartment (e.g. cytoplasm) or even if a structure is capable to move between compartments (translocation). The procedure itself consists of applying a strong laser power to consume all fluorescence (bleach) of a region of interest inside the desired compartment. Then the system continues acquiring fluorescent images. If a fluorescence is back to the bleached region it means power to that, in fact, this fluorescence recovery is due to structures outside the bleached region which are moving and are now inside this portion, proving this mobility that can be also quantified in time and intensity.



The FRET (Förster resonance energy transfer) technique is capable to detect if a structure is close of less than +-8nm to the other. To do it, the researcher needs to use known FRET pairs, two fluorochromes where the emission spectra of the donor will overlap with the excitation spectra of the acceptor. Due to this overlap, the energy of the donor emission will be capable to excite the acceptor, but only if the structures are close enough to transfer this energy. On the system itself, one extra image setup, called “FRET Image”, will be configured to excite with the donor laser line but detect on the acceptor spectra. If an image is drawn, it means that a FRET is happening. All the data within a region of interest where there’s FRET may be also quantified (“FRET efficiency”), if needed.



The conditions to have FRET: 1- Closeness between structures (Z and Y) coupled to specific fluorochromes (CFP/YFP); 2- overlap between the donor’s emission and the acceptor’s excitation to obtain a FRET Image (green zone)


What’s a laser microdissection and how does it work?


This technique makes possible to precisely isolate and excise microscopic homogeneous sub-samples from an original heterogeneous tissue section. The researcher can selectively define regions of interest, down to a single cell, cut them by a laser and collect it without contact (by gravity), to have relevant, reproducible and specific results.

The next table resumes the method of microdissection and some possible applications

 LMD1 750

As support, multiple possibilities are available. The next table show all available supports as well as the performance of all available lenses and for which application the support is optimized

 LMD2 750

Aditional information can be found at Leica website

A complete protocol guide is also downloadable



What do I have to do to have access to the platform's confocal microscope?

First, you have to contact directly the platform staff (Sandra Ormenese or Jean-Jacques Goval) or send a mail to Imaging.Giga@ulg.ac.be to book the confocal microscopy for a trial, if the experiment is not yet optimized.
Before using the system alone, you have to follow a  2 hours training. A session specially dedicated to live imaging is also available.
If the observations you have to do is urgent and you do not have the possibility to wait for the formation, you have to be accompanied by a colleague who is used to work with the confocal and is allowed to use one of our equipment.


If you do not find the answer to your question here, please contact us.