Microscopy Matters

Insights from the Advanced Imaging Center at Janelia

Lattice Light Sheet Microscopy Sample Prep

The lattice light sheet microscope (LLSM) is a powerful optical tool for observing rapidly-occurring processes in live samples. This technique offers multiple advantages over more common optical microscopy methods, like spinning-disk confocal, with improved optical sectioning, reduced photobleaching, and minimized phototoxicity. The LLSM utilizes a lattice of Bessel beams (1, 2) in order to achieve 370 nm Z resolution with 230 nm XY resolution. It also offers improved imaging depth (up to ~50 µm), which widens applicability to many models systems including D. rerio (zebrafish), C. elegans (nematode), and D. melanogaster (fruit fly). The LLSM belongs to the family of light sheet (or selective plane illumination) microscopy techniques (6), which utilize separate, orthogonally placed excitation and emission objectives (Figure 1).  This implementation in the LLSM, while giving the advantages listed above, also mechanically restricts the accessible imaging area and sample range of travel.   Due in part to that limitation, this blog will cover the most common model systems and the steps we take to mount and adapt the sample for lattice light sheet microscopy.

fig 1 w A

Figure 1. (A) Schematic depicting the orientation of objectives submerged in media bath. (B) Picture of excitation objective, detection objective, media bath, and sample holder (blue).

General Comments

Before describing mounting procedures for various model systems, I have summarized a few general points about the 5 mm coverslip sample mount and how it is utilized in the LLSM.

  • The LLSM employs a water-dipping lens on an upright configuration. Therefore, imaging requires open working space above the sample, which means there is no top cover slip.
  • The detection objective is equipped with a correction collar, which allows for the use of a wide range of media formulations while still obtaining optimal point spread functions.
  • The sample is mounted on a 5 mm coverslip, which is held in the holder by tension (Figure 2), and submerged in a media bath (Figure 1B).
  • We use 5mm round coverslips, #1 thickness, from Warner Instruments. Item number 64-0700 (CS-5R).
  • The bath does have active media exchange. Fresh media can be perfused through the bath at varying speeds, depending on need. Minimum media volume in perfusion reservoir is 40 mL. Perfused media can be thermoregulated by inline heater and thermistor.
  • Sample bath is temperature controlled via thermal conduction from circulating water channels built into the aluminum blocks.
  • Due to tight space restrictions in between the two objectives we cannot image the full ~19.6 mm2 area of the coverslip: limited to roughly +/- 1 mm in X and Y from the center.
  • Prior to use, all coverslips are cleaned in 1 M KOH and thoroughly washed with diH2O.
  • Media volume in bath is typically 10 mL.
  • Although not required, it is recommended to use genetically encoded labeling strategies (either as fusion proteins or driven by promoter of interest) to maximize the LLSM’s live-cell imaging capabilities. Dyes can also be used but the experiment must be well-planned due to likelihood of dye diffusion.

Fig 2Attached Cultured Cells

Cells that grow as an attached monolayer are the easiest and most straight-forward sample to mount. 5 mm coverslips will be prepared ahead of any visit by cleaning with potassium hydroxide (KOH), UV irradiation, and then coating with 10 µg/mL fibrinogen to improve cell attachment. The coverslips fit in a well of a 24-well culture dish. Cells can be grown on the coverslips in their typical medium (minus Phenol Red) in appropriate growth conditions. Optimal cell density should be empirically determined prior to arrival at the AIC and will be different for each experiment. When ready for imaging, the 5 mm coverslip is carefully removed from the well and slid into the holder. The holder is then affixed to the piezo stage and submerged into the pre-warmed bath containing growth media (Figure 1B).

Non-Attached Cultured Cells

Cells that grow in suspension are not as easy to mount as attachment cells, however it can be done. There will be some method variation depending on the specific cell type but generally, we first coat the cleaned coverslip in a 1:30 dilution of Cell-Tak (Corning, Product #354240). We take the suspension cells from their media and concentrate via gentle centrifugation or gravitational settling. We then take the cell suspension and carefully pipette it onto the coated coverslip and incubate for 30 min. The sample is now ready for imaging. There will likely be variability in the number of cells that remain adhered to the coverslip after submerging in the media bath so this must be considered when planning for cell density optimal for imaging.

As mentioned, this is the general protocol and modifications will be required for individual samples as multiple factors can affect stable mounting, such as sample geometry.

Danio rerio (Zebrafish)

Embryo up to 18 somite stage: Up to approximately the 18 somite stage, muscle has not yet developed so no anesthetic is necessary. Prior to imaging, we drop molten 3% agarose onto the 5 mm coverslip holder (already containing a coverslip) and allow it to solidify. We can then use a razorblade to cut a shallow triangular trough that will help orient the sample. We then carefully transfer an embryo in molten 0.5% low-melt agarose to the trough. We then use a Kimwipe to gently wick away excess agarose. Once solidified, we can proceed with imaging.

1 dpf (days post-fertilization) up to 5 dpf: The method for mounting zebrafish largely depends on the region of interest for imaging and age of the animal. For example, imaging the dorsal side requires different considerations than imaging the lateral side. Although the LLSM can image at greater depth (up to ~50 µm), one general principle is to use as little (or no) <1% agarose as possible over the imaging region. In some cases agarose must be used to stabilize the sample for non-standard orientations, but this thickness of agarose must be minimized or it will cause significant optical aberrations during imaging and can interfere with proper beam coherence at the desired penetration depth.

For imaging in the lateral orientation, we first anesthetize the animal using a Tricaine stock solution that has been diluted in Danieau’s Buffer and 0.5% low melt agarose. We also prepare a diluted solution of Tricaine to fill the media bath in order to prevent fish movement over long imaging experiments. Once the fish is immobilized, we gently pipette the fish in the molten agarose/Tricaine solution onto the 5 mm coverslip already in the holder. Using a stereo microscope we can properly orient the sample. After allowing a few minutes for the agarose to solidify, the sample is ready for imaging.

Caenorhabditis elegans (Nematode)

Embryo: A 5 mm coverslip is placed in the sample holder and coated with 5 µL of embryo glue. The glue should be given a few minutes to dry. Hermaphroditic adults are picked from their agar plate and are transferred to the coated coverslip.  Eggs are released from the adult organisms and should stick to the coverslip. Gentle manipulation can be performed using a hair loop. Proper orientation of the embryo sample is critical for imaging and can be optimized during the “pre-imaging” preparation at the AIC.

Drosophila melanogaster (Fruit Fly)

Embryo: 1 hour prior to the start of imaging, add a new agar/yeast plate to a “mating cup” in order to have fresh embryos ready. Once laid, we mount drosophila embryos using embryo glue. Approximately 5 µL of embryo glue is dropped onto a 5 mm coverslip that is already mounted in the sample holder. While the solution dries, the embryos are dechorionated by adding concentrated bleach to the agar plate (that had the yeast removed) for 1 minute. Gently but thoroughly pipette the bleach solution in order to remove all embryos from the agar surface. Using a folded Kim wipe in a funnel or fine mesh filter, pipette the bleach/embryo solution through the filter. The embryos should remain on the filter surface. Thoroughly wash with 1x PBS. Using a hair loop and a stereomicroscope, gently transfer and orient the embryos on the (now dry) 5 mm coverslip. They should securely stick to the glass and can be moved to the LLSM for imaging.



Due to wide applicability of the LLSM, we welcome any and all model systems related to life sciences. Although we cannot anticipate all possible specimens, sample preparation and mounting will be discussed in detail in pre-visit consultations. When formulating projects for the AIC, please keep in mind the details described above and how it may relate to your particular sample. As we arrive at solutions for new or unique model systems, this page will be updated.

References and Further Reading

  1. T. A. Planchon et al., Rapid three-dimensional isotropic imaging of living cells using Bessel beam plane illumination. Nat. Methods. 8, 417–23 (2011).
  2. L. Gao, L. Shao, B.-C. Chen, E. Betzig, 3D live fluorescence imaging of cellular dynamics using Bessel beam plane illumination microscopy. Nat. Protoc. 9, 1083–101 (2014).
  3. J. Durnin, J.J. Micheli Jr., J. H. Eberly, Comparison of Bessel and Gaussian Beams. Optics Letters. 13, 79-80 (1988).
  4. I.V. Majoul et al., Fast structural responses of gap junction membrane domains to AB5 toxins. PNAS. 100, 4125-33 (2013).
  5. L. Gao et al., Noninvasive imaging beyond the diffraction limit of 3D dynamics in thickly fluorescent specimens. Cell. 151, 1370-85 (2012).
  6. P. J. Keller, In vivo imaging of zebrafish embryogenesis. Methods. 62, 268-78 (2013)

Author: John Heddleston, Ph.D.

Applications Scientist

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