Calibrating Fluorescence Microscopy With 3D-Speckler (3D Fluorescence Speckle Analyzer)

Fluorescence microscopy has been widely accessible and indispensable in cell biology research. This technique enables researchers to label targets, ranging from individual entities to multiple groups, with fluorescent markers. It offers precise determinations of localization, size, and shape, along with accurate quantifications of fluorescence signal intensities. Furthermore, an ideal fluorescence microscope can achieve approximately 250 nm in lateral and 600 nm in axial resolution. Despite its integral role in these measurements, the calibration of fluorescence microscopes is often overlooked. This protocol introduces the use of 3D-Speckler (3D fluorescence speckle analyzer), a semi-automated software tool we have recently developed, for calibrating fluorescence microscopy. Calibration of fluorescence microscopy includes determining resolution limits, validating accuracy in size measurements, evaluating illumination flatness, and determining chromatic aberrations. 3D-Speckler is user-friendly and enables precise quantification of fluorescence puncta, including nanoscale 2D/3D particle size, precise locations, and intensity information. By utilizing multispectral fluorescence beads of known sizes alongside 3D-Speckler, the software can effectively calibrate imaging systems. We emphasize the importance of routine calibration for imaging systems to maintain their integrity and reproducibility, ensuring accurate quantification. This protocol provides a detailed step-by-step guide on using 3D-Speckler to calibrate imaging systems. Key features • Semi-automated particle detection. • Accurate three-dimensional measurement of fluorescent particle sizes. • High-precision three-dimensional localization of fluorescent particles. • Precision analysis of point spread function and chromatic aberration in fluorescence microscopy.


Background
Fluorescence microscopy serves as an essential tool in a variety of research fields, offering the flexibility to analyze specimens ranging from in vitro to in vivo, with resolutions spanning from micrometers to nanometers [1,2].An advantage of this technique is the ability to detect individual fluorescent targets and either simultaneously or sequentially visualize multiple fluorescent markers.Applications include the characterization of fluorescently labeled purified proteins, as well as the examination of proteins and genes within 2D/3D cell culture systems, tissues, and whole organisms.Fluorescence microscopy is utilized in various in vitro studies, such as investigating the liquid-liquid phase separation of proteins using fluorescently labeled purified proteins [3,4], examining spindle formation through droplet assays [5], and employing fluorescence resonance energy transfer (FRET) to analyze protein-protein interactions [6] and measure mechanical tension [7,8].Its utility in developmental biology extends to research across a variety of models, including plants, Drosophila, Xenopus, C. elegans, zebrafish, and mice [9].Moreover, fluorescence microscopy is a cornerstone in cell biology research, enabling the investigation of cellular processes in organisms from yeast to humans, across both 2D and 3D culture systems, as well as in living and fixed tissue samples [10,11].This technique allows researchers to accurately determine spatial locations, relative localizations, dimensions, and shapes at the nanoscale.For instance, it has been demonstrated that standard confocal microscopy can achieve <5 nm precision in determining 2D/3D protein architecture within cells using a dual-color fluorescence approach [12,13].Furthermore, fluorescence microscopy excels in quantifying protein copy numbers and assessing relative protein concentrations with high accuracy [14][15][16].The resolution of fluorescence microscopy is constrained by the point spread function (PSF), which represents the 3D diffraction pattern of light emitted from an infinitely small point source [17].An ideal fluorescence microscope, equipped with a high numerical aperture (NA) objective lens (NA > 1.4) and a high-resolution camera, can achieve resolutions of approximately 250 nm laterally and 600 nm axially [18,19].However, the maximum resolution is also contingent on the signal-to-noise ratio (SNR) of the specimens.These resolution limits can be enhanced without altering the optical setup through expansion microscopy (ExM), which physically expands specimens without changing the optics setup, or through post-imaging processing techniques, such as deconvolution [18,20,21].Additionally, ongoing developments in super-resolution microscopy significantly improve resolutions compared to conventional fluorescence microscopes, enabling the further detailed study of nano-scale cellular structures [1,[22][23][24].
All fluorescence microscopes have the potential to achieve their theoretical resolution maximum and quantitative accuracy.However, this potential heavily relies on the components and the conditions under which microscopy is conducted.To understand the current conditions of the imaging system, it is critical to routinely perform calibrations.We have recently developed the 3D-Speckler software, which enables precise quantification of fluorescence puncta in both biological and non-biological samples [18].Utilizing fluorescence beads of known sizes in conjunction with 3D-Speckler allows for the comprehensive evaluation of resolution limits, size quantification accuracy, chromatic aberration determination, and the flatness of illumination produced by imaging systems.The aim of this article is to present a comprehensive guide on utilizing the 3D-Speckler software for calibrating imaging systems.This guide is intended for a broad audience interested in fluorescence microscopy, rather than experts in optics and mathematics.Consequently, this protocol minimizes the explanation of the coding and mathematical principles behind 3D-Speckler.For detailed information, please refer to our original manuscript [18].Briefly, 3D-Speckler identifies fluorescence particles primarily based on their relative signal intensities against the background and determines particle sizes through a full-width-at-half-maximum (FWHM) calculation based on a 2D or 3D Gaussian profile fitted to the intensity profile.The intensity measurements for fluorescence beads can be performed either without background correction or with a local background correction for each bead [14].3D-Speckler can determine the center of fluorophores with ~2 nm accuracy, which allows it to accurately identify chromatic aberrations, an unavoidable image distortion that occurs between different wavelengths, by measuring the distance between the centers of fluorophores with different wavelengths within a single bead.This process is designed to evaluate their current performance and ensure their integrity, accuracy, and reproducibility in quantification tasks.

A. Obtaining images of fluorescent beads
For calibration with 3D-Speckler, we recommend utilizing fluorescent beads of at least two different sizes: one smaller than the point spread function (PSF) of your imaging system (less than 200 nm) and one larger (for example, 250 nm, 500 nm, or 1 μm).Smaller beads will be used to determine the lateral resolution limit of your system, which is equivalent to its PSF.Larger beads will be used to evaluate whether your system can accurately measure the size of beads larger than the PSF.In this protocol, we used TetraSpeck beads of 100 and 500 nm.3D-Speckler can be used for both 2D and 3D images of fluorescence beads obtained from any fluorescence microscope.Additionally, 3D images can be utilized for 2D calibration (see details in our original manuscript [18]).Axial size measurement and 3D chromatic aberration correction require 3D stack images.1.Power on the microscope system and select the objectives you plan to use for calibration.2. Place a slide with TetraSpeck beads (100 nm) on the microscope stage.3. Locate a field of view (FOV) that contains TetraSpeck beads.We recommend finding a FOV containing fluorescence beads that are evenly distributed and not aggregated or too crowded.This helps minimize the difficulty and errors in identifying individual beads and taking measurements.Then, finely adjust the focus, the camera's exposure time, and the light source's power for optimal imaging (see General note 5). 4. (Optional) Set the z-range to encompass the entire depth of the beads, ensuring that the interval between each step is less than 200 nm (see General note 6). 4 Published: Aug 20, 2024

5.
Acquire images at the wavelengths you wish to calibrate.It is recommended to capture images at different wavelengths at the same z-plane for chromatic aberration measurements.6. Repeat steps A2-A5 at several different locations on the slide to ensure thorough calibration (see General note 7).7. Using a slide with larger TetraSpeck beads (for example, 500 nm), repeat steps A2-A6 (Figure 1).

3D-Speckler interface features
Figure 2 displays the primary user interface of 3D-Speckler.A detailed description of each module is listed below.If users select an image that is single-wavelength, 3D-Speckler will prompt users to decide whether to add more wavelength images (Figure 3) (see General note 13).
Figure 3. Selecting image wavelengths.Upon selecting an image file, it will be displayed.You will then need to choose the wavelengths and review both lateral and axial scales.
e.The main user interface will launch upon clicking choose.After completing this step, the 3D-Speckler base interface will appear (Figure 4   l.Review and manage detected particles: To remove unwanted particles, first click the Select button found beneath the Fit Gaussians button.Then, within their bounding boxes, double-click on the particles you wish to eliminate (Figure 7).Once you have reviewed the detected objects, click on Save Channel Results to save the data for the current channel.Ensure you save your data before moving to the next channel; otherwise, unsaved measurement results will be lost.Then, specify the desired location to save the files.Upon doing so, multiple plots will be displayed, followed by a confirmation message from 3D-Speckler indicating "Data Export Completed Successfully."

Reviewing results
a. 3D-Speckler will create a new folder that includes an Excel file, MATLAB images for each channel, and a subfolder containing field-dependent plots of FWHM and signal intensities.The MATLAB images of beads will feature assigned numbers for each detected bead, corresponding to the numbering in the Excel file.b.To review and obtain the 2D and 3D size of TetraSpeck beads, open the Excel file (Figure 9).3D-Speckler offers three distinct methods for measuring FWHM: true (tFWHM), interpolated (iFWHM), and Gaussian FWHM (gFWHM).The differences lie in the methods used to obtain the intensity profile of detected particles for measuring FWHM.Detailed descriptions of these measurements can be found in our original article [18].Briefly, gFWHM and iFWHM use Gaussian and interpolation fitting methods, respectively, whereas tFWHM does not employ mathematical fitting methods.We recommend using tFWHM for lateral measurements and iFWHM for axial measurements [18].The average lateral size (tFWHM) can be determined by averaging the values of the blue columns in the Excel file (Column Q and R in an Excel file in 3D measurements).Similarly, the axial size (iFWHM) can be calculated by averaging the values in the black column (Column V in an Excel file in 3D measurements) (see General note 18).The lateral and axial sizes of beads smaller than the PSF represent the resolution limit of your system (measured PSF in your system).You can compare and evaluate these values against the theoretical resolution limit of your system.The measured size of beads larger than the PSF should be equal to or close to their actual size.If not, it may indicate that the bead size you used is still smaller than the PSF of your system or suggest potential defects in your system.  1.
Table 1.Definition of the abbreviations in Figure 9. Explanation of the abbreviations in the output Excel file.

Abbreviation Explanation
Gauss_X, _Y, -Z x, y, and z coordinates of the center position of detected particles.
Theta_X, _Y, _Z x, y, and z angles based on the optimum center of detected particles, as described in the original 3D-Speckler manuscript.STD_X, _Y, _Z Standard of deviation of Gaussian fitting.

GaussBase
The base signal intensity of Gaussian fitting.If users select local background correction option, this value is used for local background correction.

GaussPeak
The peak signal intensity of Gaussian fitting.tFWHM_X, _Y, _Z tFWHM values of each detected particle in x, y, and z-axis.iFWHM_X, _Y, _Z iFWHM values of each detected particle in x, y, and z-axis.gFWHM_X, _Y, _Z gFWHM values of each detected particle in x, y, and z-axis.

ResNorm
Normalized residual value: how well the objects fit the 2D/3D Gaussian model.

Validation of protocol
This protocol or parts of it has been used and validated in the following research article • Jonathan Loi, Xiaofei, Qu, Aussie Suzuki (2023).Semi-automated 3D fluorescence speckle analyzer (3D-Speckler) for microscope calibration and nanoscale measurement.J Cell Biol.(Figures 1-8).

General notes
11. Executing 3D-Speckler by clicking Run does not require changing the working directory to include the "bfmatlab" folder.However, to use the "Open3DSpeckler" command for operation, it is necessary to change the working directory.12. 3D-Speckler automatically reads and categorizes images into different channels.If only one single is detected, it allows users to import data for more channels.13. 3D-Speckler automatically detects the lateral and axial resolution by using the image metadata.If these resolutions are not automatically filled in or the numbers have errors, users must manually input or correct the scaling values.14.The 3D-Speckler interface automatically updates based on the selected analysis options and workflow.Particle Analysis is the default analysis method for analyzing general fluorescent particles.15.For advanced settings, 3D-Speckler provides a Local Background Correction option for signal intensity and Gaussian fitting measurements.Since background signals are not uniform across the FOV, local background correction emerges as one of the most accurate measurement methods [14,15,18].To utilize this feature, users need to optimize parameters for further analysis.The default is set at 0.15 (which is 15% larger in all dimensions than the bounding box used for measurements in each detected particle to obtain local background signals).For most analyses involving TetraSpeck beads, this default setting proves effective.16.For an additional optimization step, by clicking on multi-thresholding, users can select a threshold to commit an image.3D-Speckler will then remove those objects, allowing users to set another threshold until all desired objects are accurately detected.Conclude by clicking on Finish Thresholding.17.The Distance Metrics option offers 2D or 3D distance measurements between all detected beads, both within a single channel and across different channels [18].This feature is not necessary for microscope calibration.18. 3D-Speckler offers three different measurement methods for FWHM.Please see the details in the original article [18].19.This measurement boasts a precision of less than 5 nm [18].20.3D-Speckler provides correction for chromatic aberration and outputs the corrected images, as detailed in the original article [18].

Figure 1 .
Figure 1.Example of a TetraSpeck bead image.The 500 nm TetraSpeck beads were imaged using a 60× objective lens.The full field of view (FOV) image is displayed on the left.On the right, the XY and XZ views of a single 500 nm bead from the left image are presented.Visualization of XY and XZ views using Imaris software (Andor).

Figure 4 .
Figure 4. 3D-Speckler main interface.After completing wavelength selection, this interface will be displayed.You will need to select either Particle Analysis or Local Chromatic Aberration Correction for downstream analysis.

Figure 5 .
Figure 5. Thresholding bar.It pops up when you click Start Analysis.Users can adjust the threshold by sliding the bar to the right or left.

Figure 6 .
Figure 6. Background correction buffer.This option becomes available after you click on Fit Gaussians.If you wish to perform background correction, please select the size of the bounding box for background measurements.Choosing 1 will double the dimensions of the bounding boxes used for quantification (see General note 15).

Figure 7 .
Figure 7. Review fitting results.After fitting, particle locations will be shown in blue, not green.Click the select button located under the Fit Gaussians button, and then double-click on particles within their bounding box to remove them.

Figure 8 .
Figure 8. Matching and aligning detected particles across all channels.The Match & Align Pts function enables the selection of corresponding particles across different channels, eliminating those that do not match.The white arrow indicates a particle unmatched in the Blue channel, resulting in its removal from quantification in all channels, as shown by the Red arrows.

10 Published: Aug 20, 2024 Figure 9 .
Figure 9. Quantification results.Example results of the green channel.The output is in Excel file format.Abbreviations' definitions are listed in Table1.
c. 3D-Speckler measures integrated and maximum signal intensities for each bead [the red columns (O and P) in Figure9].Using this information, 3D-Speckler can generate field dependence plots that illustrate variations in the FWHM measurements and intensity across the FOV of an imaging setup.Users will have the option to export these plots.If chosen, 3D-Speckler will save the plots as .figfiles in the Field Dependence Plots folder.These plots can then be analyzed to identify areas within the field of view that provide the most accurate measurements (Figure10).

Figure 10 .
Figure 10.Visualization of field dependency plots.Surface plots of integrated intensity and maximum intensity variations illustrate changes across the imaging field of view (FOV).The light source in fluorescence microscopy is typically brightest at the center of illumination and gradually becomes dimmer toward the edges.

Figure 11 .
Figure 11.Aberration between different channels.The Excel file contains the worksheets named "aberration_wavelength".These worksheets include aberration in x, y, and z-axis and 3D distance measurements (µm scale) (see General note 19-20).