Light-sheet microscopy (Huisken et al.) is a versatile 3D imaging tool with advantages such as fast imaging, minimal phototoxicity and photobleaching, and the ability to view millimeters into tissue and from different angles. As such, it is ideal for live imaging of large samples in 3D and is widely used e.g. to describe early embryonic development in zebrafish (Keller et al., Mongera et al.), to visualize gastrulation events in Drosophila embryos (Amat et al.), and to characterize developing organoids (de Medeiros et al.).
Why I use light-sheet microscopy: I built my own light-sheet microscope to characterize how mammalian embryo models (gastruloids) grow and develop. Especially, we are interested in how the first organs form. To fully utilize my volumetric imaging tool, I also study Drosophila embryos, zebrafish larvae, and cell spheroids in collaboration with various groups at UiO.
Workflow: I collect image stacks at various depths into the tissue, and from different angles. To enhance the images, I deconvolve them using Generic Deconvolution, and I use Matlab and Huygens to obtain 3d representations from the deconvolved data.
Optics and laser illumination: Our light-sheet microscope (Figure 1) is a stripped-down version of the Open-SPIM platform (Pitrone et al.). The illumination laser (Cobolt 6 series, Hübner Photonics) has laser heads with four different wavelength (λ = 375, 488 561, 647 nm), enabling us to visualize the gastruloid nuclei (λ =488 nm) and membrane (λ =375 nm), as well as the mesoderm and endoderm germ layers (λ =488 nm, 561nm).
A fiber optic cable safely guides the laser light to a beam head with a built-in collimator. To minimize photobleaching of the specimen, the beam passes through a neutral density filter (ND507A, ThorLabs), and to form the light sheet, it passes through a cylindrical lens (ACY254-050-A, f = 50 mm, ThorLabs), two mirrors (POLARIS-K1, ThorLabs) and an adjustable slit (VA100/M, ThorLabs). Two spherical lenses (AC127-050-A-ML, f = 50 mm, ThorLabs; AC127-025-A-ML, f = 25 mm, ThorLabs; spaced 75mm apart) serve to collimate the light, and a water dipping lens (UMPLFLN 10XW, 10X/0.5, Olympus) focuses the light on to the specimen, giving an in-plane spatial resolution of 1.5 μm in x and 2.1 μm in y, and an out-of-plane (z) resolution of 6.7 μm (as measured by the full width half maximum of the point spread function).
Image acquisition: To minimize optical distortions (Weber et al.), the specimen rests on an agarose plug (concentration: 2%) within a pre-rinsed fluorinated ethylene propylene (FEP) tube (BOLA, ID=0.8 mm, OD=1.6 mm) with refractive index closely matching water (n ≈1.33) inside of a water filled acrylic viewing chamber (22 mm ×22 mm×37 mm). A movable stage (USB-4D-Stage, Picard Industries; resolution: 1.5μm) holds the FEP tube, and interfaces with a lab computer to allow the operator to rotate and to translate the sample along the x, y and z axes. Finally, the imaging system consists of a water immersion objective (UMPLFLN 20XW, 20X/0.5, Olympus) and an emission filter (OD6 ULTRA Quad-Bandpass, Alluxa), which sit in front of a light sensitive camera (Andor Zyla 5.5, Andor). A lab computer controls the stage and camera via Micro-Manager (Edelstein et al.), while Cobolt Monitor interfaces with the same computer to control the laser.
Live imaging: Motion capture of cells in 3D
To describe how cells multiply, specialize and move, we perform live imaging and track them in 3D. We create physiological conditions by controlling the temperature, pH and oxygen level.
Temperature control
To control the temperature, we circulate pre-heated water at physiological temperature through channels in the viewing/incubation chamber, see Figure 2.
References
Amat, F., Lemon, W., Mossing, D. P., McDole, K., Wan, Y., Branson, K., ... & Keller, P. J. (2014). Fast, accurate reconstruction of cell lineages from large-scale fluorescence microscopy data. Nature methods, 11(9), 951-958.
Edelstein, A., Amodaj, N., Hoover, K., Vale, R., & Stuurman, N. (2010). Computer control of microscopes using ?Manager. Current protocols in molecular biology, 92(1), 14-20.
Huisken, J., Swoger, J., Del Bene, F., Wittbrodt, J., & Stelzer, E. H. (2004). Optical sectioning deep inside live embryos by selective plane illumination microscopy. Science, 305(5686), 1007-1009.
Keller, P. J., Schmidt, A. D., Wittbrodt, J., & Stelzer, E. H. (2008). Reconstruction of zebrafish early embryonic development by scanned light sheet microscopy. Science, 322(5904), 1065-1069.
de Medeiros, G., Ortiz, R., Strnad, P., Boni, A., Moos, F., Repina, N., ... & Liberali, P. (2022). Multiscale light-sheet organoid imaging framework. Nature Communications, 13(1), 4864.
Mongera, A., Rowghanian, P., Gustafson, H. J., Shelton, E., Kealhofer, D. A., Carn, E. K., ... & Campàs, O. (2018). A fluid-to-solid jamming transition underlies vertebrate body axis elongation. Nature, 561(7723), 401-405.
Pitrone, P. G., Schindelin, J., Stuyvenberg, L., Preibisch, S., Weber, M., Eliceiri, K. W., ... & Tomancak, P. (2013). OpenSPIM: an open-access light-sheet microscopy platform. Nature methods, 10(7), 598-599.
Weber, M., Mickoleit, M., & Huisken, J. (2014). Multilayer mounting for long-term light sheet microscopy of zebrafish. JoVE (Journal of Visualized Experiments), (84), e51119.