Tutorial: guidance for quantitative confocal microscopy

Abstract

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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Fig. 1: Quantitative confocal microscopy example.
Fig. 2: Principles of confocal microscopy.
Fig. 3: Effect of wavelength on resolution.
Fig. 4: Effect of mounting medium on confocal images of fixed cells.
Fig. 5: Comparing inter-related key instrument performance parameters of different microscopy techniques and configurations.
Fig. 6: Objective lens comparisons.
Fig. 7: The effects of photobleaching during widefield observation.
Fig. 8: Configuring confocal detection channels.
Fig. 9: Unexpected instrumentation issues cause uncertainty in intensity measurements.
Fig. 10: Sampling and statistics.

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Acknowledgements

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.

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Contributions

No section of this manuscript was untouched by all five authors. J.J. drafted the outline, assembled the team and wrote the Introduction, ‘Removing bias’, ‘Troubleshooting instrumentation issues’ and ‘Analyzing and presenting quantitative images’. He also generated Figs. 1, 2, 6, 7, 8 and 9; Table 1; Box 3; and Supplementary Figs. 1 and 2 and contributed to general editing. C.M.B. analyzed adhesion dynamics data, generated the statistics figure (Fig. 10), wrote the Statistics section of the manuscript and contributed to ‘General considerations for preparing samples for quantitative fluorescence microscopy’, ‘Preparing fixed cells and tissues’ and ‘Preparing live cells’ and significantly to general editing of the manuscript. G.D.W. worked on ‘Choosing the right microscope’ and ‘Setting up the microscope’, performed general editing of the manuscript and generated Fig. 5, the images for Box 2 and Supplementary Video 1. K.I.A. worked on ‘Choosing the right microscope’ and ‘Planning your experiment’ and performed general editing of the manuscript. A.J.N. worked on ‘General considerations for preparing samples for quantitative fluorescence microscopy’ and ‘Setting up the microscope’, contributed extensively to general editing and generated Figs. 3 and 4 and Boxes 1 and 2.

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Correspondence to James Jonkman.

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The authors declare no competing interests.

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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.

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Integrated supplementary information

Supplementary Fig. 1 How objective magnification affects the brightness of a CLSM image.

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.

Supplementary Fig. 2 Focus drift.

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 information

41596_2020_313_MOESM3_ESM.mov

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.

Supplementary Information

Supplementary Figs. 1 and 2.

Reporting Summary

Supplementary Video 1

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

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