TITLE: Ensuring accurate co-registration measurement for quality control of Single Point Confocal Laser Scanning Microscopes - V1
AUTHORS: Aurelien Dauphin, Maria Azevedo, Aidan Boyce, Fabrice Cordelières, Herlinde De Keersmaecker, Frank Eismann, Orestis Faklaris, Hans Fried, David Grunwald, Christian Kukat, Pascal Lorentz, Tobie Martens, Ileana Micu, Roland Nitschke, John Oreopoulos, Allister DA Pattison, Alex L Payne-Dwyer, Vincent VTG Schoonderwoert, Christian Schulz, Maria Smedh, Sathya Srinivasan, Nicolas Stifani, Kees van der Oord, Nicolaas van der Voort, Marlies Verschuuren, Luis-Francisco Acevedo-Hueso, Judith Lacoste

[DESCRIPTION]
Chromatic aberration is a major source of spatial inaccuracy in multi-color fluorescence microscopy, arising from the inability of optical systems to bring different wavelengths to the same focal position. These wavelength-dependent shifts can originate from objective lens imperfections, refractive index variations, or misalignment of optical components, and often manifest as lateral or axial displacements between fluorescence channels. Accurate co-registration of multi-color images is therefore essential for reliable interpretation of spatial relationships in biological specimens. 
This protocol provides a streamlined and quantitative method to assess co-registration accuracy in laser-scanning confocal microscopes, following the principles of ISO 21073. While the ISO-recommended procedure is laborious and time-consuming, our approach offers a faster and more user-friendly alternative while remaining compatible with ISO specifications. Using 1-µm multi-color beads, the protocol enables precise measurement of channel alignment, facilitating routine performance monitoring and high-quality multi-color imaging.

[BEFORE_START]
Chromatic aberration refers to possible artifacts caused by a wavelength-dependency of the imaging properties of any fixed optical beam path of the microscope system (as a result of optical design of the system, the manufacturing tolerances of the system, and the current state of alignment of the optical components). 

Co-registration accuracy more generally refers to the ability of the system to co-localize dyes of different wavelengths during a particular experimental set-up. Depending on the experimental set-up and also the system architecture of the system to be tested, one might achieve different results. This could occur if the system architecture or experimental set-up force the system to use different beam paths for the experiment. The required switching of beam paths introduces an additional complication for the co-registration on top of the chromatic aberrations.

1) Prior knowledge of the microscope architecture is important in order to be able to set-up experiments with minimal mechanical switching movement for highest accuracy results. If the experiment requires mechanical switching of optical components, all available channel calibrations/adjustments should be performed successfully before any quality control measurement (for example pinhole alignment and beam splitters).

2) If the system allows for chromatic corrections (for example, different collimators for different excitation sources) or adjustments (software-based corrections), all corrections and adjustments have to be optimized prior to measurements and kept stable during measurements.

3) Objective lenses have different levels of correction for chromatic aberrations. Therefore they will differ in their performance with respect to spectral bandwidth within a given aberration budget and in their maximum aberration over a given spectral range. For example, objectives rated (according to ISO19012-2)  as semiapochromats will typically have a smaller spectral bandwidth as well as larger residual aberrations within the visible wavelength range than apochromats. Semiapochromats are typically marked with a "FL" in their designation, like "Fluor", "FLUOTAR" or "Neofluar". Apochromats always contain "Apo" in their designation. Objectives which exceed the specifications of an apochromat are labelled as, for example, 
"APO CS2", "Apo Lambda", "C-Apochromat" or "XAPO" to indicate an even higher level of chromatic correction.

4) The system and the specimen should also be in steady temperature state. To confirm a steady state is reached, recording the temperature during measurements close to the sample is recommended. Capturing this information in the metadata is recommended.

5) When using oil immersion objective, the immersion oil to be used should be clear to the eye, devoid of crystals, dirt etc. 
In addition, be cautious to use only the specified oil for immersion objectives since small differences (in particular variations of the Abbe number) may influence the co-registration accuracy significantly.  Capturing this information in the metadata is recommended (manufacturer, catalog number, lot number, refractive index, expiration date (if any) etc.).

[GUIDELINES]
This protocol focuses on measuring the co-registration accuracy of a single point confocal laser scanning microscope in x, y, and z for each fluorescence color available in this type of system. We define the following:

I. Sample Preparation.
II. Pre-requisites
III. Image Acquisition
IV. Data Analysis

[STEPS]
SECTION: Placing the sample on the microscope
5. Clean the slides with only 70% ethanol prior to imaging.  


 

SECTION: Beads positioning on the coverslip
4. Using tweezers, remove a clean coverslip from its 100% ethanol storage bath. Carefully remove excess liquid with lens tissue and let the coverslip air dry while resting on another clean lens tissue surface.
SECTION: Acquisition settings
6. Prior knowledge of the microscope architecture is essential for setting up experiments with minimal mechanical switching movement for the highest accuracy results. If the experiment requires mechanical switching of optical components, all available channel calibrations/adjustments should be performed successfully before any quality control measurements (pinhole alignment and beam splitters).  If the system allows for chromatic corrections (for example, different collimators for different excitation sources) or adjustments (software-based corrections), all corrections and adjustments have to be optimized before measurements and kept stable during measurements.
SECTION: Coverslip preparation
1. Prepare coverslips, grade #1.5H (170 ± 5 μm) thickness (according to ISO8255-1). Immerse the coverslips in a 1M HCl solution and shake gently for 30 min at room temperature. 30 min 

SECTION: Coverslip preparation
1.1. Rinse the coverslips twice with distilled water and immerse them in 100% ethanol for storage (up to several months). 

SECTION: Bead Selection
1.2. Beads with different diameters and fluorescent properties can be used for this protocol. Here, beads co-emitting blue (360/430 nm), green (505/515 nm), orange (560/580 nm), and dark red (660/680 nm) fluorescence with 1 µm diameter were used. The choice is based on tests (see Supplementary Information, Section 11). 

  



SECTION: Beads Dilution
3. Beads must be diluted so their density is high enough to find the focus quickly on the microscope, but also low enough to prevent bead aggregation and overcrowding. A final dilution of around 100 beads per μl is recommended (once on the coverslip, the density is around 1 bead per 100 μm²). 
SECTION: Beads Dilution
3.1. Centrifuge the tube containing the bead stock solution in a benchtop centrifuge to collect all the liquid at the bottom. 

SECTION: Beads Dilution
3.3. Dilute the stock beads in distilled water at 1:2500. If sub-dilutions are made, vortex the solution for 1 min at each step.


 


SECTION: Beads Dilution
3.2. Vortex the stock solution tube for 3 minand then sonicate it in a water bath for 2 min to dissociate possible aggregates.

SECTION: Beads positioning on the coverslip
4.2. Vortex the final diluted bead solution for 3 min.

SECTION: Beads positioning on the coverslip
4.3. Add ten small drops (~1 μl) of the bead suspension onto the coverslip. Carefully place the first drop in the center of the coverslip and place the other drops around (total amount 10μl - See Figure 1). Let the drops dry in the dark at room temperature. 180 min



 



SECTION: Beads positioning on the coverslip
4.4. Place a drop of mounting media (e.g., ProLong‱ Gold Antifade Mountant, Thermo Fisher Scientific, P10144, or ProLong‱  Glass Antifade Mountant, Thermo Fisher Scientific, P36980) on the microscope slide (e.g., 10 µL drop of mounting media for 24x24mm coverslips). Position the coverslip upside-down on the microscope slide and remove any excess mounting medium from around the edges of the coverslip using filter paper, ensuring that the coverslip sits flat against the larger glass slide. Follow the mounting medium specification time for curing.


  

  

 
       

SECTION: Beads positioning on the coverslip
4.1. Obtain a standard 1 mm thick microscope slide and clean it with 95% ethanol. Carefully remove excess liquid with lens tissue and let the slide air dry on another clean lens tissue surface.
SECTION: Beads positioning on the coverslip
4.5. Once the mounting medium has cured, or if it is non-hardening, the coverslip edges can be sealed to the slide with silicone Picodent Eco-Sil. 


  


 


 



 



SECTION: Placing the sample on the microscope
5.3. Choose the immersion medium according to the objective's specification and the temperature used for imaging - oil usually has a refractive index of ne=1.518 at 23°C.
 
 

SECTION: Placing the sample on the microscope
5.2. If available, set the objective lens correction collar to the correct position (0.17 mm and correct temperature setting if present).


 


SECTION: Placing the sample on the microscope
5.1. Clean the microscope objective lens.
SECTION: Acquisition settings
6.1. Set the pinhole diameter to 1 Airy Unit (AU), calculated for 520 nm emission. This value is usually computed automatically by the acquisition software; however, the user may need to manually adjust the reference wavelength or emission channel settings to ensure the calculation is based on 520 nm. Use the same pinhole diameter for all four acquisition channels.


 


SECTION: Data Analysis
7. Co-registration analysis can be performed by first splitting the acquired Z-scan image dataset into its respective channels, when necessary for the software package used, and subsequently localizing the center point coordinates of the beads with sub-pixel accuracy in each channel. Once these sub-pixel coordinates are known for a bead, the 3-dimensional distances between the measured center points in each channel can be calculated in µm. The separation distances for any two-channel pairs are used as a metric of the given microscope configuration co-registration for that channel pair. A separation distance of zero indicates perfect co-registration for a given channel pair. In contrast, a separation distance greater than zero, which is more realistic and often measured, means that the microscope configuration exhibits some level of mis-registration in that channel pair. What constitutes an acceptable degree of channel pair separation distance will be addressed in the discussion of this protocol.
SECTION: Data Analysis
7.2. Before proceeding with the co-registration analysis, it is recommended to first check the XZ or YZ projection views of the datasets for any obvious signs of non-spherical or asymmetric aberration, which might indicate an issue with the system optics or alignment. 

SECTION: Before starting
2. The bead size, fluorescence specifications, manufacturer, catalogue number, lot number, and expiration date (if any) should be recorded. The same applies to all reagents used throughout this protocol.
SECTION: Acquisition settings
6.2. If differential interference contrast (DIC) optics (polarizers, analyzers, prisms, etc.) are available, remove them from the light path.
SECTION: Acquisition settings
6.3. 

For image acquisition, choose a pixel size about 2.3x smaller than the characteristic spatial resolution of the imaging system. 

The theoretical estimative of the Full Width at Half Maximum (FWHM) of the Airy unit (1 AU) of the Point Spread Function (PSF) was used to calculate the lateral resolution (ResL):

  

and the axial resolution (ResA): 

  

λex refers to the excitation wavelength. As a reference for all channels, we set λex to 488 nm, which corresponds to the excitation wavelength used to generate our emission reference at 520 nm. Here, n denotes the refractive index of the imaging medium, and NA is the numerical aperture of the objective lens. Reference: MetroloJ_QC gitHub: https://github.com/MontpellierRessourcesImagerie/MetroloJ_QC)
 

 

 


 


 




SECTION: Acquisition settings
6.4. Acquire a volume of at least 5 µm in x, y, and z directions for the TetraSpeck 1 µm beads, with an image at least every 200 nm.
SECTION: Acquisition settings
6.5. Acquire all the available fluorescence colors using suitable lasers and filters/detectors.


 



 

SECTION: Acquisition settings
6.6. Use 4x line averaging and a pixel dwell time of around 1 µs.
SECTION: Acquisition settings
6.7. Use the highest bit depth available on your detector (12-bit, 16-bit). When setting the detector offset, avoid zero values in the image and ensure the signal-to-background ratio (SBR) is equal to or greater than five by adjusting the laser power as required while using the above-recommended dwell times and averaging settings.

For example, if using a detector with a 16-bit dynamic range, adjust these acquisition parameters such that the brightest pixels in the beads exhibit an intensity of 4000 grayscale levels or arbitrary digital units (ADUs). Ensure you have no saturation in the images. 


 



 

SECTION: Acquisition settings
6.8. Image as many beads as possible to minimize statistical variances that occur naturally. It is recommended that at least 3 beads on at least 3 FOV be acquired.
SECTION: Acquisition settings
6.9. Store the images in the original, uncompressed data format.
SECTION: Acquisition settings
6.10. Save your datasets in the proprietary file format provided by your microscope manufacturer, to ensure all necessary metadata is retained. Additionally, record the parameters outlined in the list available below.

  

SECTION: Placing the sample on the microscope
5.4. Allow the system to warm up to prevent excessive drift and laser fluctuation during the acquisition.


 

SECTION: Placing the sample on the microscope
5.5. Focus on the beads using a filter set matching their fluorescence properties (e.g., use the orange emission filter to focus the beads, as the TetraSpeck beads are brightest at this emission wavelength). The focal plane is right at the coverslip-mountant interface.


 



SECTION: Placing the sample on the microscope
5.6. Focus on one bead (choose a bead with intensity from the medium-bright population of beads visible in the sample) and move the stage to center this bead in the middle of the field of view (FOV).


 

SECTION: Data Analysis
7.1. 
We advise the use of MetroloJ_QC, as it is an open-source plug-in of Fiji, and provides many interesting metrics.

However, various software options are available for performing co-registration analysis (Table 1).

 


 



SECTION: Data Analysis
7.3. For microscopes used for colocalization experiments, it is recommended to run this protocol at least after installation and maintenance. 
SECTION: Supplementary Information
11. Beads Selection

Different bead sizes can be used to assess co-registration accuracy in microscopy systems. Multi-color TetraSpeck beads of sizes 0.1, 0.2, 0.5, 1, and 4 µm are commercially available. We evaluated the performance of Tetraspeck 0.2, 1, and 4 µm beads side by side across 12 microscopes, with testing conducted by 11 members of the Quarep-LiMi WG4. Data analysis was performed using MetroloJ_QC (version 1.3.1.2), a plugin for Fiji/ImageJ software. 


The green channel was used as the reference to determine the distance to the far-red (Ch0), red (Ch1), and blue (Ch3) channels. Our results indicate that 1 and 4 µm TetraSpeck beads have comparable performance, providing consistent measurements in the XY and Z-axis (Figures 3, 4, and 5). In contrast, Tetraspeck 0.200 µm beads exhibit higher variability and dispersion in measurements across all channels, particularly along the Z-axis (Figures 3B, 4B, and 5B), likely due to their lower achievable signal levels compared to the larger bead sizes. Additionally, 0.2 µm beads may be more prone to forming clumps that can be difficult to identify during image acquisition. 


 


 


 

For the 0.2 µm beads, two different sampling metrics were tested: 0.016 x 0.016 x 0.040 µm and 0.070 x 0.070 x 0.200 µm (Figures 6, 7, and 8). The results showed that smaller voxel sizes yield more precise and robust distance measurements to the reference channel. The SBR was comparable between the two voxel sizes, indicating that the primary difference was in the number of voxels and, consequently, the number of photons.


 

 

 

The co-registration distance relative to the microscope’s theoretical resolution was calculated for all bead sizes. As shown in Figures 9A, 9B, and 9C, for the 4 µm and 1 µm beads, the ratios are below the system’s theoretical resolution, indicating that they are performing correctly with respect to co-registration. For the 0.2 µm beads, the data show greater dispersion; however, the majority of the analyzed beads fall within the reference ratio of 1. The same data are also represented in Figure 9D, where the distance ratio between the blue–green (B–G) channel and the red–green (G–R) channel is shown for all beads, and the Figure 9E, where the distance ratio between the Far red–green (G-FR) channel and the red–green (G–R) channel is shown for all beads. The green region indicates values with acceptable co-registration. In this representation, the beads with a ratio greater than 1 are easy to track, and it becomes clearer that the 0.2 µm beads are indeed problematic when analyzing the blue–green and the far red-green distances.  


 

Despite the higher standard deviation observed with the Tetraspeck 0.2 µm beads, the average distances calculated to the reference channel were similar across all three bead sizes and the two voxel sizes.

Several parameters influence the ability to localize a bead, including its size, brightness, and the microscope's point spread function (PSF). The higher standard deviation observed with Tetraspeck 0.2 µm beads may result from their relatively weak fluorescence signal, as they contain fewer fluorescent molecules, which can lead to a low SBR and makes localization analysis more challenging. In contrast, due to their larger volume, 1 and 4 µm beads produce stronger fluorescence signals that improve the SBR, facilitating more precise localization analysis.

Although smaller beads, such as the Tetraspeck 0.2 µm, enable higher localization precision, we recommend using Tetraspeck 1 µm beads for routine co-registration analysis, as they yield higher SBR and provide more stable and reliable results, as our findings indicate. Therefore, 1 µm beads are better suited for routinely assessing the co-registration performance of microscopy systems, particularly in high-resolution systems where strong signal intensity is crucial, such as confocal laser scanning microscopes.
Of note, if localization precision is a key factor for a specific application, smaller diameter beads, such as the Tetraspeck 0.2 µm beads, should be used, guaranteeing that a sufficiently high SNR can be achieved. If a system is equipped only with low magnification objectives, the Tetraspeck 4 µm would be preferable.


 

 

 
 MetroloJ_QC gitHub: https://github.com/MontpellierRessourcesImagerie/MetroloJ_QC)
SECTION: Data Analysis
7.4. 
 

SECTION: Data Analysis
10. If your images exhibit any misregistration, you can refer to the troubleshooting section for guidance.





SECTION: Data Analysis
8. Here are the main steps to follow if you are using MetroloJ_QC plugin (version 1.3.1.2) on ImageJ :

Launch ImageJ (or Fiji) and open the MetroloJ_QC toolbar (Plugins > MetroloJ_QC)
Open your 3D multichannel stack (single 1 µm bead, 4 channels)
Click on the Co-registration Tool icon

 

SECTION: Data Analysis
8.1. In the Microscope Acquisition Parameters: 
Choose the microscope type (CLSM), the objective specification, the pinhole size, and the excitation wavelengths.
SECTION: Data Analysis
8.2. In the Bead Detection Options: 
Choose a thresholding method ("Minimum" is advised for noisy images);
Select “Centroid” as the center detection method;
Enable “Discard bead if more than one particle is thresholded". Set the background annulus size (for signal-to-background calculation);
Return to the main window, select "PDF report" and "Excel spreadsheet" in the "File Save Options", and run the analysis.
SECTION: Data Analysis
9. The tool will compute three useful metrics : 
SECTION: Data Analysis
9.1. Shifts in X, Y, Z between all channel pairs: These represent the pixel distances between the bead centers in each channel pair along the X, Y, and Z axes. They quantify how misaligned the channels are in each spatial dimension.
SECTION: Data Analysis
9.2. Center-to-center distances: This refers to the 3D Euclidean distance between the centers of beads in two different channels. It combines the X, Y, and Z shifts into a single value and is calculated as:

  
SECTION: Data Analysis
9.4. Profile view images (XY, XZ, YZ) and outlines of the thresholded bead will be shown for visual inspection.
SECTION: Data Analysis
9.3. Distance ratios relative to the reference resolution : This is the ratio of the measured center-to-center distance to the theoretical resolution of the microscope in the angle of the shift. 
It helps assess how significant the shift is relative to the system’s resolution.
A ratio < 1 indicates acceptable co-registration; a ratio > 1 suggests potential misalignment beyond the system’s resolving capability