When used appropriately, a confocal fluorescence microscope is an excellent tool for making quantitative measurements in cells and tissues. The confocal microscope’s ability to block out-of-focus light and thereby perform optical sectioning through a specimen allows the researcher to quantify fluorescence with very high spatial precision. However, generating meaningful data using confocal microscopy requires careful planning and a thorough understanding of the technique. In this tutorial, the researcher is guided through all aspects of acquiring quantitative confocal microscopy images, including optimizing sample preparation for fixed and live cells, choosing the most suitable microscope for a given application and configuring the microscope parameters. Suggestions are offered for planning unbiased and rigorous confocal microscope experiments. Common pitfalls such as photobleaching and cross-talk are addressed, as well as several troubling instrumentation problems that may prevent the acquisition of quantitative data. Finally, guidelines for analyzing and presenting confocal images in a way that maintains the quantitative nature of the data are presented, and statistical analysis is discussed. A visual summary of this tutorial is available as a poster (https://doi.org/10.1038/s41596-020-0307-7).
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J.J. thanks the AOMF staff for helpful discussions, Courtney McIntosh for the images in Supplementary Fig. 1, and the Princess Margaret Foundation for ongoing financial support of the AOMF. G.D.W. thanks A*STAR and the National Research Foundation’s Shared Infrastructure Support Grant for continued support of the A*STAR Microscopy Platform and John Common for samples (Box 2). C.M.B. acknowledges Alex Kiepas (McGill University), who collected the adhesion dynamics data for the statistics section of the paper including Fig. 10, and the ABIF for general support and access to the Diskovery spinning disk TIRF microscope for collecting the adhesion dynamics data. K.I.A. thanks the Francis Crick Institute for their CALM support, and facility colleagues for helpful discussion. A.J.N. thanks the Rockefeller University for its continued support of the Frits and Rita Markus Bio-Imaging Resource Center (BIRC), the Sohn Conference Foundation for funding the Leica SP8 confocal microscope used to generate Figs. 3 and 4 and the facility staff and users for stimulating discussions.
The authors declare no competing interests.
Peer review information Nature Protocols thanks Gary Laevsky, Timo Zimmermann and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Guidance for Quantitative Confocal Microscopy (https://doi.org/10.1038/s41596-020-0307-7): This poster is a visual summary of this paper.
Molecular Expressions Optical Microscopy Primer (https://micro.magnet.fsu.edu/primer/): Developed by Michael W. Davidson and his team at The Florida State University, this website hosts a vast amount of knowledge on all aspects of optical microscopy from the physics of light and color and the anatomy of a basic transmitted-light microscope to advanced techniques such as FRET and TIRF.
iBiology Microscopy Series (https://www.ibiology.org/online-biology-courses/microscopy-series/): Directed by Ron Vale, Nico Stuurman and Kurt Thorn, this 72-video series (and growing!) starts with the basics of optical microscopy and concludes with some of the latest techniques, such as super-resolution microscopy. Their Short Microscopy series of just 14 videos may be more manageable for some.
iBiology BioImage Analysis Course (https://www.ibiology.org/online-biology-courses/bioimage-analysis-course/): Created by Anne Carpenter (CellProfiler project lead) and Kevin Eliceiri (ImageJ project lead), this series focuses on general steps of image processing as well as introductions to CellProfiler and ImageJ specifically.
Microscope Vendor websites: Many microscope vendors have educational material on their websites. Some of them license sections from Molecular Expressions, but supplemented with their own microscopes’ unique attributes. Others have written their own material from scratch.
The Confocal Microscopy Listserv (https://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy): With an active subscriber base of nearly 4,000 participants and an archive that goes back to 1991, the Confocal Listserv is a valuable resource for confocal and related questions. Registration (free) is required.
Microforum (https://forum.microlist.org/): This new web forum led by Jennifer Waters and Tally Lambert was established in 2019. Its focus is on hardware, acquisition and specimen-related aspects of scientific imaging, particularly (but not limited to) the theory and practical use of optical microscopes and detectors, fluorescent proteins and probes and specimen preparation.
Integrated supplementary information
a, Schematic showing illumination and light collection for 40×/1.4 NA and 63×/1.4 NA objective lenses. The same objective is used for both illumination and collection in an epifluorescence geometry, but for clarity, the beampath has been unfolded. The incoming laser beam is focused to a small point in the specimen plane: the PSF. The shape of the PSF is determined by the NA of the objective. The intensity of the PSF depends on the size of the back aperture of the objective, which changes with the magnification of the lens. The incoming laser beam overfills the back aperture of the objective, with the result that the back aperture crops the outer rays of the laser beam, reducing the intensity of the incoming laser beam accordingly. For example, when switching from a 40×/1.4 NA objective to a 63×/1.4 NA objective, the aperture area is reduced by a factor of 402/632 = 0.40. Hence, we would expect a CLSM image through the 63× objective to be ~40% of the intensity compared to using the 40× objective, if the NA and quality of the lenses are identical. Does this mean that a 40×/1.4 NA objective is more sensitive than a 63×/1.4 NA objective because of the increased brightness through the 40× lens? No. The increased brightness for the 40× lens stems from the fact that more of the laser beam passes through the aperture and hits the sample: this change in intensity by means of a physical aperture is no more helpful than changing the laser power in the software. On the other hand, a higher NA is always helpful for confocal microscopy since the collection efficiency of the objective increases with the NA2 independent of magnification. b, CLSM image of a mouse kidney slide labeled with Alexa Fluor 488-WGA (Molecular Probes Prepared Slide #3) taken with a 40×/1.4 NA objective. c, CLSM image of the same field of view as b, taken with a 63×/1.4 NA objective using the same imaging parameters as b. The mean intensity of the image is reduced by ~36% in the 63× lens compared to the 40× lens. However, the power of the laser beam, which was measured using an oil immersion–compatible power meter (Thorlabs PM400 console with S170C sensor), was also reduced by 34%. This demonstrates that the dominant effect of changing magnification is to crop out a portion of the laser beam, for which one could easily compensate by increasing the laser power accordingly. The scale bar is 10 μm.
Focus drift affects most microscope stands for 2–3 h after turning the microscope on, even when no incubators are used. A thin, fixed fluorescently labeled slide (Fluocells Prepared Slide #1, Molecular Probes) was placed on the stage of several confocal microscopes that had been turned off overnight (≥12 h). An optimized image was captured, and the focus position was recorded 10 min after turning power on to the instrument. For subsequent time points, the focus was adjusted manually so that the new image exactly matched the first image of the time series (the saturation LUT was helpful for evaluating when the images matched). There were no microscope incubators installed on these microscopes. a, Focus drift measured three times on the same Leica SP8 equipped with STED superresolution and a Super Galvo Z-stage demonstrates that the stand should be turned on 2–3 h before beginning confocal acquisition (particularly if STED is employed, as the acquisition times tend to be longer than regular confocal imaging). The Leica DMi8 microscope stand’s closed-loop focus feedback was enabled. b, Focus drift on a similar Leica SP8, both with and without the Super Galvo Z-stage, and both with and without the DMi8 closed-loop focus enabled. c, Focus drift on four other microscopes, demonstrating that the problem is not limited to any particular brand but is widespread.
Supplementary Figs. 1 and 2.
DIC complements fluorescence for live-cell confocal microscopy. Live-cell confocal fluorescence (left) and DIC (right) timelapse imaging of keratinocytes expressing Keratin5-GFP. DIC can produce sharp images of cell and organelle boundaries without the need for labeling with additional fluorophores.
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Jonkman, J., Brown, C.M., Wright, G.D. et al. Tutorial: guidance for quantitative confocal microscopy. Nat Protoc 15, 1585–1611 (2020). https://doi.org/10.1038/s41596-020-0313-9