Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Summary

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Abstract

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Introduction

Ensuring minimal phototoxicity is a major requirement for imaging live embryos with high spatiotemporal resolution for long periods. Over the last decade, light sheet microscopy has become the methodology of choice to meet this requirement^{1,2,3,4,5,6,7}. Briefly, in this technique that was first used in 2004 to capture developmental processes⁸, two aligned thin sheets of laser pass through the embryo from opposing ends, illuminating only the plane of interest. A detection objective is placed orthogonally, and then the emitted fluorescent light from all illuminated points in the sample is collected simultaneously. A 3D image is then obtained by sequentially moving the embryo through the static light sheet.

In addition, in a specific form of this methodology, termed multiview light sheet microscopy, the samples can be suspended in a polymer tube that can be rotated using a rotor, enabling imaging of the same embryo from multiple angles^9,10,11. Following imaging, the images from multiple angles are fused based on registration markers, which are typically globular fluorescent markers within the embryo (e.g., nuclei) or in the tube (e.g., fluorescent beads). Multiview imaging and fusion significantly improve the axial resolution, providing isotropic resolution across all three dimensions¹². While this is a big advantage, a major challenge of multiview methodology is sample mounting, where embryos have to be mounted and kept in place in the tubes during the entire time course of imaging.

For performing multiview imaging, to keep the embryos in place and prevent movement while imaging, embryos can be embedded in agarose. However, this often leads to detrimental growth and development, particularly for early-stage zebrafish embryos¹³, the model system that is discussed here. A second mounting strategy is to use a thin tube that is only slightly bigger than the diameter of the embryo, where the embryo can be pulled into the tube along with the embryo medium, followed by closing the bottom of the tube with an agarose plug¹⁴. In this method, because the tube is filled with embryo medium, registration markers such as fluorescent beads cannot be used for the fusion of the different views, and registration is therefore reliant on markers within the embryo. In general, beads act as better registration markers as the signal of the markers within the embryo degrades upon moving deeper into the sample owing to both illumination and detection limitations of any microscope.

Thus, a third approach, which will be detailed here and used previously^{5,13,14,15,16}, is imaging early zebrafish embryos with an intact chorion and filling the tube with a minimal percentage of agarose, which contains beads as registration markers. In this scenario, because manual intervention for positioning embryos within a chorion is not possible, this study provides statistics on the default orientation early zebrafish embryos fall into, particularly focusing on 70% epiboly and bud stages. It then discusses the optimal number of views required for imaging early-stage embryos at cellular-scale resolution and details the process of fusion using BigStitcher, a FIJI-based plugin^10,17,18. Together, this protocol, which uses a 20x/1 NA objective, aims to facilitate zebrafish embryologists in using multiview light sheet systems for imaging embryos with nuclei and membrane markers from gastrulation to early somite stages.

Protocol

The zebrafish maintenance and experimental procedures used in this study were approved by the institutional animal ethics committee, vide Reference TIFR/IAEC/2023-1 and TIFR/IAEC/2023-5. Embryos obtained by crossing heterozygous fish expressing Tg(actb2:GFP-Hsa.UTRN)¹⁹ were injected with H2A-mCherry mRNA (30 pg) at the one-cell stage. H2A-mCherry mRNA was synthesized using the pCS2+ H2A-mCherry plasmid (a gift from the Oates lab, EPFL) by in vitro transcription. Embryos expressing both markers, referred to as Utr-GFP and H2A-mCherry, respectively, in the rest of the protocol, were imaged at the 70% epiboly and bud stages. The details of the reagents and equipment used in the study are listed in the Table of Materials.

1. Sample preparation for multiview imaging

1. Preparation of FEP/PTFE tubes
   1. Clean FEP/PTFE tubes following previously described procedures¹⁴.
   2. After cleaning, cut the tubes into lengths of 2-2.5 cm as per requirement and store them in 2 mL microcentrifuge tubes containing double distilled water. Tubes can be stored in this manner for about a month.
   3. Before sample preparation, heat the tubes at 70-75 °C for approximately 25 min to straighten the tubes. Avoid overcrowding in each microcentrifuge tube to prevent clumping, ensuring sufficient space for effective tube straightening.
      NOTE: Wear gloves during the entire tube handling procedure, as any fingerprint/dust particle left on the tubes may interfere with imaging.
2. Preparation of agarose
   1. Prepare E3 buffer 50x stock solutions as follows:
      1. Stock 1 - 3.252 g of Na₂HPO₄, 0.285 g of KH₂PO₄, 11.933 g of NaCl, 0.477 g of KCl in 1 L of deionized water
      2. Stock 2 - 2.426 g of CaCl₂, 4.067 g of MgSO₄ in 1 L of deionized water.
   2. To make 1x E3 buffer, add 20 mL each of stock 1 and 2 solutions in 960 mL deionized water.
      ​NOTE: Do not add methylene blue in the E3 buffer used for imaging as it causes scattering of light.
   3. Dissolve low-melting point agarose in 1x E3 by heating at 70-75 °C on a heat block with stirring until the solution becomes clear with no clumps or crystals.
   4. For multiview imaging, add commercially available fluorescent beads (see Table of Materials) to the agarose solution, which enables image registration during analysis. Sonicate the beads at room temperature with 40 kHz frequency for 20-30 min in a water bath ultrasonicator to disaggregate the beads.
   5. After sonication, add 1 µL of beads to 10 mL of agarose solution in a 15 mL tube and vortex the tube well to ensure uniform dispersion of the beads.
      NOTE: The volume of bead solution to be added to agarose depends on the stock solution and the manufacturer. Try a range of volumes and choose a concentration where the beads appear well dispersed when imaged in the light sheet system. Too many beads can aggregate despite sonication and vortexing, while too few beads can affect image registration.
   6. Keep the agarose tube with added beads in a closed water bath maintained at 37 °C for at least 30 min before sample preparation. This ensures that the temperature of the agarose comes down from 70 °C to 37 °C.
3. Sample preparation
   1. Set up the fishes in pairs with dividers in the evening before the day of imaging. Remove the dividers for about 15 min and collect embryos using standard procedure²⁰.
   2. Once the embryos reach the stage of interest for sample preparation, pour the entire 10 mL of agarose (with beads) that is kept at 37 °C onto a 6 cm Petri dish.
   3. Wait for about 2-3 min to bring the temperature lower than 33 °C before transferring 10 to 15 embryos to the agarose solution.
      NOTE: This is an important step for preventing any possible heat shock to embryos transferred to the agarose.
   4. Ensure that as little E3 buffer as possible is added to the agarose solution while transferring embryos. Swirl the Petri dish so that the little buffer that would have been transferred gets well dispersed.
   5. Wear gloves for the rest of the sample preparation procedure.
   6. Take a 200 µL micropipette with an appropriate pipette tip and insert a cleaned straight tube taken out of the microcentrifuge tube to the pipette tip as shown in Figure 1A,B.
   7. Aspirate a little agarose into the tube, followed by 2-3 embryos using the micropipette (Figure 1C,D). Ensure that the embryos are close to the bottom of the tube (Figure 1E), which will become important while setting up imaging. Also, ensure that there is a little agarose in between the different embryos (Figure 1E) so that beads in this region can be used as registration markers.
   8. Without releasing the pressure from the pipette, detach the tube from the pipette tip and place it on the lid of the petri dish filled with E3 for the agarose to solidify (Figure 1F).
   9. Repeat steps 1.3.6 to 1.3.8 until all embryos in the Petri dish are transferred to the tubes.
   10. Once the agarose has solidified (which can take 5-10 min and can be confirmed by checking the agarose in the Petri dish), transfer the tubes to a 2 mL microcentrifuge tube filled with E3 (Figure 1G).
   11. Store the tubes with the embryos in a 28 °C or 33 °C incubator, as required, before proceeding to multiview imaging.

2. Multiview imaging

NOTE: This step presents a general procedure for multiview imaging of zebrafish embryos at their early development stages. The method detailed below can be easily adapted to any multiview light sheet microscopy system.

1. Locating the embryo
   NOTE: Fill the sample chamber of the microscope with the embryo medium²⁰ and set the temperature of the sample chamber to the temperature of interest at least 10 min before imaging.
   1. Assemble the sample holder as previously described²¹, however, with one modification. Since imaging will be performed through the tube, insert the tube with embryos directly into a capillary of the right diameter, such that it is held in place without pushing the agarose out and disturbing the embryos at the bottom (Figure 1H).
      NOTE: Since the sample chamber has a defined height, ensuring that the embryos are located towards the bottom of the tube during sample preparation (as mentioned in step 1.3.7) would enable positioning the embryo in the field of view without hitting the floor of the chamber.
   2. Insert the sample holder into the multiview system and use the x- and y- controls to bring the embryo to the center of the field of view.
   3. Use the z control of the specimen navigator to then bring the embryo into focus.
   4. At this point, the orientation of the embryo within the chorion will be clearly visible, and if it is not satisfactory, insert a new tube and choose the embryo with an orientation of interest.
2. Aligning the light sheets
   1. Once the embryo is in position, set up all the experimental settings - lasers, light path, filter, beam splitter, and camera.
   2. While setting up the acquisition parameters, if the software provides an option for pivot scan, select it. A pivot scan will reduce the shadowing effect emerging from the light sheet as it encounters any structures or opaque regions in the sample.
   3. Start live scanning of the sample. Set up the bit size, laser power, and desired zoom, which will determine the thickness of the light sheet and, hence, the axial resolution.
   4. To align the light sheets, first switch to single-sided illumination and change the settings of the two light sheets (left and right) sequentially. Digitally zoom into a region of the sample where clear structures of interest are visible.
      NOTE: In this study, either a nuclear (H2A-mCherry) or a membrane (Utr-GFP) marker was used to align the sheets. Changing the light sheet settings in the software is recommended so that the sharpest contrast of signal is achieved in the region of interest.
   5. Once the two light sheets are individually optimized to achieve the best signal, switch on both light sheets. Perform a second round of alignment and ensure that flickering is minimal, which indicates a fairly good alignment of the two sheets.
   6. Once alignment is done, activate the option of automatically fusing the images generated by the two sheets, if allowed by the software. Alternatively, one can first coarsely align the light sheets by focusing on beads around the sample and then fine-tune the alignment using information from the sample.
      NOTE: While imaging with two fluorophores (such as nuclear and membrane markers), the optimal alignment is usually marginally different for the two structures. In this scenario, choose a setting based on constraints on downstream processing. For example, if cell boundaries are to be segmented, which is relatively more processing intensive, align the light sheets based on the membrane marker.
3. Setting up multiview imaging
   1. After aligning the light sheets, activate the options for performing a z-stack as well as multiview imaging.
   2. Start live scanning and navigate using the specimen navigator to select the first view. In this view, set up the slice interval, followed by the first and last slices of the z-stack. Add this position in the multiview dialogue box, which takes note of the position and z-stack information for this view.
   3. Move to the next view by changing the angle in the specimen navigator. Set up the z-stack for the second view and add the information in the multiview dialogue box.
   4. Repeat the same for additional views.
      NOTE: Ensure that a certain minimum number of slices are imaged in each view so that there is enough overlap between the different views, which will eventually aid in registering the views during processing. The minimum number of slices will depend on the zoom factor as well as on the number of angles chosen for imaging.
   5. After setting all the views, save this information into a text file (named 'embryo-positions', for example), which will contain z-stack information, (x, y, z) coordinates of the tube as well as specifications of the angle for each view. After saving, clear all positions from the multiview dialogue box.
   6. Since imaging is done with an intact chorion, which is relatively large, it is not possible to view the beads in the tube with the same settings. Therefore, bead information has to be acquired from a different position in the tube. To do this, translate to a different 'y' coordinate away from the embryo where beads are visible. Add this position to the multiview dialogue box and save this position in a text file (named 'beads-y', for example).
      NOTE: Image the beads as close to the embryo sample as possible in order to minimize the effects of possible differences in tube curvature at the two positions. Therefore, if multiple embryos are mounted in the tube, it is important to leave a little agarose in between the tubes to image beads (as mentioned in step 1.3.7).
   7. Go to the folder where the text files are saved and duplicate the 'embryo-positions' file to a new file named 'beads-positions'. Replace the y-coordinate for all the views in the 'beads-positions' file with the y-coordinate from the 'beads-y' file. This will ensure that beads are imaged with the same number of views, z-stacks, and x-coordinates but at a different y-position in the tube.
   8. Return to the imaging software and load the 'bead-positions' file in the software. If the time series option is activated, select one cycle and start the experiment. Save bead images as 'beads', which will be used for registering the different views during image processing.
   9. After taking the bead images, clear the positions and load the initial 'embryo-positions' file in the software. Set the desirable number of cycles with a suitable time interval and start the experiment.

3. Multiview image analysis

NOTE: For fusing the multiview images, a FIJI plugin, BigStitcher, which is the latest version of the Multiview Reconstruction plugin, is utilised^10,17,18. The plugin can be installed by adding the BigStitcher plugin in the 'Manage Update Sites' function, which can be accessed in the 'Update' option under the 'Help' menu. Once installed, the plugin will appear under the 'Plugins' menu. The broad steps involved in fusion are as follows: (1) Define .xml/.h5 file pairs for both the beads and the embryos; (2) Register all the views with the beads file; (3) Extract Point Spread Function (PSF) for the beads, which can be used for deconvolution (Figure 2A); (4) Transfer the registration and PSF information from the beads file to the embryo file and begin multiview deconvolution. Most of these steps have been previously described in detail²¹, and here, the steps that are differently processed are described.

1. Defining a .xml dataset
   1. Define a new dataset for both the beads and the embryo as 'beads.xml' and 'embryo.xml', respectively, as previously described²¹.
2. Detection and registration using interest points
   1. Following the successful export of the beads.xml file, select the views that are required for registration in the 'Multiview Explorer' dialogue box (Figure 2B). Right-click and choose Detect Interest Points. Proceed with the steps as described²¹.
   2. After detecting interest points, select all the views, right-click, and choose Register using Interest Points. Follow the protocol as described²¹.
   3. Carefully check if the bead registration has worked successfully. Upon successful registration, overlapping beads from different views get superimposed (Figure 2C). To check for how precisely registration has worked, select two consecutive views, go to the overlapping region, and toggle between the two views to observe if the beads imaged from different angles are superimposed. Repeat this for every consecutive view.
   4. If the overlap is not precise, retry registration by relaxing the required significance for registration and/or the acceptable error of overlap (termed 'RANSAC error' in the 'Registration' dialogue box).
   5. Following successful registration, click on Save in the 'Multiview Explorer' window to save the updated .xml file.
      NOTE: Save the log file as well, as it contains further detailed information on the efficiency of registration. To translate the registration information from the beads to the embryo, follow the steps described below.
   6. Open the bead.xml file in a text editor of choice and copy the entire block under "ViewRegistrations".
   7. Open the embryo.xml and replace the "ViewRegistrations" block with the block copied from the beads file. If there are multiple channels, replace the registration information for each channel as above. The registration information can be transferred either manually or by using the custom-written MATLAB code that can be downloaded here: https://github.com/sundar07/Multiview_analysis
   8. Open the embryo.xml file in "BigStitcher" and carefully check if the registration has worked successfully for the embryo. Repeat the same as done for the beads, by checking the overlap of structures of interest in every two consecutive views.
      NOTE: Occasionally, embryo registration may not be as desirable despite the beads getting registered perfectly. This is possible if there are slight changes in tube curvature from the position where the beads are imaged to the embryo position. In addition, there could be subtle movements of the embryo between views as it is freely floating within the chorion. In this case, perform a second round of registration using a nuclear marker in the embryo.
   9. To do this, open the embryo.xml file in "BigStitcher", right-click on all the views that have the nuclear marker, and choose Detect Interest Points.
   10. Rename the interest points as 'nuclei' and proceed with the steps as performed for beads.
   11. While setting 'Difference of Gaussian' parameters, ensure that most, if not all, of the nuclei are detected and that there is no ectopic nuclear detection. Next, click on Done.
   12. Following this, register these views by choosing Register using Interest points and opt for the Precise descriptor based (translation-invariant) option. Ensure that the 'compare all views and interest points' option is selected and use 'nuclei' as interest points. Use the option for fixing the first view and do not map back.
   13. For registration, use an affine model with rigid regularization and the default parameters in the plugin.
   14. Re-check the success of registration by comparing every two consecutive views.
   15. Following successful registration, click on Save in the 'Multiview Explorer' window to save the updated .xml file.
   16. Open the embryo.xml file in a text editor of choice and copy the registration information from the nuclear markers to the other channels.
3. Point spread function extraction and assignment
   1. To perform multiview deconvolution, extract the PSF of the imaging system from the registered beads dataset and apply it to the embryo file.
   2. To get this information, select all the views in the bead file and then right-click and choose Point Spread Functions and Extract options.
   3. In the dialogue box that appears, ensure that the Use Corresponding Interest points and Remove min intensity projections from PSF are checked, proceed with the default PSF sizes, and click on OK.
      NOTE: If the PSF extraction is successful for all views, the log file will display 'Extracted n/n PSFs'
   4. Following this, resave the .xml file. A ticked checkbox in the PSF column of the 'Multiview Explorer' dialogue box will appear, and a 'psf' folder will be generated in the respective folder with all extracted PSFs.
   5. Open the embryo.xml file and assign the PSF for each view separately. Right-click "a view" → click on Point Spread Functions → choose Assign → select Advanced, followed by Assign new PSF to all selected views. Click on Browse, go to the .xml file path, and open the psf folder.
   6. Choose the matching PSF with the corresponding ID in the selected view and click on OK, following which the PSF checkbox will appear ticked in the 'Multiview Explorer' window.
   7. Repeat the process for all other views.
4. Multiview fusion and deconvolution
   1. After the point spread function has been assigned for all the views, right-click and choose Multiview Deconvolution.
   2. Select the bounding box as currently selected views. For this work, the default OSEM acceleration and number of iterations work well.
   3. Downsample the images as required if faster computation is desired or if CPU memory is limiting.
      NOTE: If the required RAM exceeds the existing memory, a warning error message in red color will pop up at the bottom of the window. Do not start the deconvolution if this warning pops up, as the plugin will stall and stop responding at some point during the processing.
   4. To assess the progress of deconvolution, check the log file, which will display results every 5 iterations.
   5. To increase the computational speed, perform multiview deconvolution in GPU if previously installed.
      NOTE: At the end of this process, a fused image window will pop up, which can be saved as a tiff file.

Results

Orienting the sample in a precise manner is a vital part of efficiently using a microscopy set-up. However, manually orienting samples is often not possible when using a multiview light sheet system, given the requirement for preparing the samples in a tube. Therefore, to check if there are stereotypical positions that embryos take up within the chorion, zebrafish embryos were imaged at 70% epiboly (about 7 h post-fertilization (hpf)), since time-lapse imaging from gastrulation to early somite stages was the focus of thi...

Discussion

Positioning an embryo in the right orientation to image the region of interest is one of the rate-limiting steps that often results in a failed microscopy session for a user. This is more so in a multiview light sheet microscope where manual manipulation of the orientation is difficult as the samples are embedded within a tube. To aid in this process, this study reports the statistics of various positions a zebrafish embryo takes up between 70% epiboly and early somite stages within a chorion when the polymer tube with e...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Agarose, low gelling temperature	Sigma-Aldrich	A9414
Calcium Chloride dihydrate	Sigma-Aldrich	12022
FIJI			Version: ImageJ 1.54f
Latex beads, carboxylate-modified polystyrene, fluorescent red, 0.5 μm mean particle size, aqueous suspension	Sigma-Aldrich	L3280
Magnesium sulfate heptahydrate	Sigma-Aldrich	M2773
mMESSAGE mMACHINE SP6 Transcription kit	ThermoFischer Scientific	AM1340	For in vitro transccription of H2A-mCherry plasmid
Potassium Chhloride	Sigma-Aldrich	P9541
Potassium phosphate monobasic	Sigma-Aldrich	P0662
PTFE Sleeving AWG 15L - 1.58 mm ID x 0.15 mm Wall +/-0.05	Adtech Innovations in Fluoroplastics	STW15	PTFE tubes
Sodium Chloride	Sigma-Aldrich	S3014
Sodium phosphate dibasic	Sigma-Aldrich	71640
Ultrasonic Cleaner	Labman	LMUC3	Ultrasonicator
Zeiss LightSheet 7 System	Zeiss