Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

1. Assembling the diSPIM steel imaging chamber with poly-L-lysine coated coverslip

NOTE: The steps below are all required to optimize and automate lineage analysis of nematode embryos by StarryNite/AceTree. Several options (indicated by as such) may be omitted for experiments that do require tracing of C. elegans cell lineages.

1. Draw a small rectangle (2mm x 5mm) in the center of a clean rectangular coverslip (no. 1.5, 24 mm x 50 mm) with a sharpie (or similar pen).
2. Flip the coverslip over to the unmarked side and place 10 μL of poly-L-lysine (cat. no. P1524) over the marked rectangle.
   NOTE: Make a working concentration of 1 mg/mL poly-L-lysine dissolved in filtered water (or equivalent). For long-term storage, prepare 5–10 mL aliquots and store at −20 °C. Once thawed, the aliquots can be stored at room temperature (23 °C) for 3–4 weeks.
3. Allow the poly-L-lysine to coat the coverslip for 5 minutes (Figure 1A). Poly-L-lysine is used for coating glass cover slips where the embryos will be mounted, allowing the embryo eggshell to stick firmly to the coverslip, even when the objectives are immersed in the imaging medium.
4. Place the poly-L-lysine coated coverslip in the bottom half of the steel imaging chamber.
5. Place the top half of the steel imaging chamber onto the bottom half with the coverslip and tighten with the four screws associated with the chamber. Check visually from the side to make sure the top half is evenly seated in the bottom half (Figure 1B).
6. Fill the chamber with 7–8 mL of M9 buffer¹⁹, an isotonic buffer which helps prevent early stage embryos from succumbing to abnormal osmotic pressure. 1-cell, and even 2-cell embryos are osmotically sensitive and can develop abnormally if not in this isotonic buffer. Embryos also tend to arrest at the 3-fold stage if M9 is substituted with water.

2. Preparing C. elegans embryo samples for mounting

NOTE: Approximately 18 hours prior to imaging, five young (1 day since final molt) adults and ten larval stage 4 (L4) C. elegans are picked to a nematode growth medium (NGM) agar plate seeded with E. coli strain OP50. Platinum wire picks are used to move larvae and young adult C. elegans without harming the animal¹⁹.

1. Prepare a 1% methyl cellulose (cat. no. H7509-25G) solution in M9 buffer.
   NOTE: Methyl cellulose must be stirred in warm M9 until it is dissolved. Once prepared, this solution can be stored at room temperature.
2. Add 500 µL of 1% methyl cellulose-in-M9 solution into the depression of a concave microscope slide. This viscous buffer will be used in two occasions: 1) when harvesting early embryos by dissection of adult worms and 2) when washing late-stage embryos picke­­­­d directly from an NGM plate.
   NOTE: Methyl cellulose is used to prevent embryos from sticking to the microscope slide.
3. For imaging late stage embryos, pick laid C. elegans embryos (preferably from an NGM plate with young adults present) by using an eyelash pick, and move the embryos to the 1% methyl cellulose on the concave microscope slide. The eyelash pick helps reduce force and thus minimize stress or damage to the embryos during handling. The procedure to make an eyelash pick is covered by Hart²⁰.
4. With a second eyelash pick (in the opposite hand), gently tap both eyelashes together to suspend the embryos in the methyl cellulose.
5. Optional: If planning to lineage embryonic cells with StarryNite, one must mount 1-cell to 4-cell embryos. To do so, first select young adults from an NGM plate and move them into the M9-methyl cellulose solution on the concave microscope slide by using a platinum wire pick.
6. Optional: With the sharpened tips of hypodermic needles (no. 18G x 1 ½), slice the animal transversely at the mid body to release 1-cell to 4-cell embryos.

3. The Mouth Pipette: Assembling aspirator tube with microcapillary pipette

NOTE: We use an aspirator tube with a hand pulled microcapillary pipette inserted into the rubber gasket of the tube. This allows us to transfer embryos from the dissection slide to the Poly-L-lysine-coated surface in the buffer-filled imaging chamber.

1. Manually pull the microcapillary pipette over an open flame to create two halves with stretched tips.
2. Take one half of the microcapillary pipette and insert the blunt end into the rubber gasket of the aspirator tube (Figure 1C). Set the other half of microcapillary pipette aside for use later (if needed).
3. With the assembled aspirator tube fitted with microcapillary pipette, gently break the tip of the microcapillary pipette and create an opening that will fit about 1–2 two embryos (from here on this instrument is called a “mouth pipette").

4. Mounting C. elegans embryos on Poly-L-lysine coated coverslip

1. With the aspirator mouthpiece held gently between the teeth, pre-fill the microcapillary pipette with 10–15 µL of M9 Buffer and then gently suck several embryos from the concave slide into the capillary.
2. Transfer the embryos to the steel imaging chamber filled with M9 Buffer, positioning the capillary tip so that embryos will fall into the central rectangle of the coverslip.
3. Avoiding injury to the embryos, gently move them with an eyelash pick or use the mouth pipette to position the embryos vertically, to orient the embryos so that the long axis of the embryo is perpendicular to the long axis of the coverslip (Figure 1B inset, bottom panel).
   NOTE: Positioning the embryo in this orientation minimizes the number of slices to image, thereby reducing light dosage and data processing time while improving acquisition speed.
4. Place the steel imaging chamber into the sample holder on microscope stage (Figure 1D).

5. Assembly, software setup and laser optimization for embryonic imaging using the diSPIM

1. See step-by-step instructions on how to assemble the entire fiber-coupled diSPIM from commercially available parts in Kumar et al.^10,11 and at http://www.dispim.org. A video-protocol of how to assemble the diSPIM is also available on the ASI website (http://www.asiimaging.com).
   NOTE: Instrument setup for this protocol is identical to Kumar et al.^10,11 diSPIM, which makes use of 40x 0.8NA water immersion lenses for imaging. The only difference between the setup in this protocol and Kumar et al.^10,11 is the addition of a dichroic mirror (splitting at 560 nm) and red and green bandpass filters inside an imaging splitting device (model A12801-01) installed on both diSPIM imaging arms. The addition of image splitting optics allows simultaneous capture of images from two distinct fluorophores – excited by 561 nm and 488 nm lasers – by separating the emission bands onto two halves of the same camera chip.
2. After instrument assembly, check the optical alignment of the diSPIM before imaging.
   NOTE: To properly align the diSPIM see https://youtu.be/qnOrg30NNuE, and for hardware information, http://dispim.org/hardware/objectives and http://www.asiimaging.com.
3. Use the open-source platform Micro-Manager (https://micro-manager.org/)²¹, which has been optimized for operating Light Sheet Microscopes for high-throughput cellular imaging²². We recommend use of the ASI diSPIM plugin for multi-position acquisition, which allows simultaneous imaging of up to 30 embryos as described²³.
4. With Micro-Manager open, set laser intensities to ~179 µW (0.5) for 488 nm and ~79 µW (0.25) for 561 nm (Figure 2A, red rectangle).
   NOTE: These are recommended settings for long-term imaging of C. elegans embryos using 1-minute intervals. During long-term dual-color imaging, the 561 nm laser is used to image nuclei (mCherry::Histone) until embryos are at bean stage, at which point the 488 nm laser is then turn on to also image the GFP-labeled neurons. These imaging conditions are optimized for minimizing phototoxicity and ensuring survival and hatching of the embryos while enabling prolonged (12–14 hours) continuous acquisition of neurodevelopmental and lineaging data.
5. In Micro-Manager, choose menu Plugins > Device Control > ASI diSPIM to open the ASI diSPIM window (Figure 2B). Choose the Acquisition Tab. In this tab's Data Saving Settings section (green rectangle), Volume Settings section (blue square), and Slice Settings section (orange square), ensure that each parameter is set as shown in Figure 2B.
   NOTE: Our image analysis software CytoSHOW is adapted to work with other optional output data formats such as bulk-concatenated OME-TIFF file series and TIFF-stack file series created post-acquisition through use of an export function built into Micro-Manager. Typically, the single-time-point-stack collated OME-TIFF file data format is used because it allows real-time viewing and processing of image volume as soon as the raw data are acquired.

6. Optimized autofocus parameters for long-term imaging of C. elegans embryos

1. Set Micro-Manager autofocus parameters to the settings optimized for long-term lineage-quality diSPIM-imaging of C. elegans embryos. In the ASI diSPIM window, click the Autofocus Tab (Figure 2C). In the General Autofocus Options section (black square) specify parameters precisely as shown. Note that the Autofocus channel (red square) should specify your nuclear channel fluorescence channel in lineaging experiments.
   NOTE: If the max offset is greater than 5 µm, images tend to drift out of focus.
2. Click, Plugins > Acquisition Tools > Pattern Overlay.
3. In the Pattern Overlay window, click Show Grid.
4. In the ASI diSPIM window, click the Navigation tab.
5. Click check boxes for Beam and Sheet of Path A or B then click Live. Image acquisition begins. A Live view window opens. Select the auto-focus analysis region of the embryo by drawing a box around the embryo on the selected channel from 6.1.
   NOTE: We typically capture 420 time points for 10 embryos per imaging session. The raw data per image session are typically 1.7TB, while deconvolved and StarryNite-processed data are 1.4TB (see Steps 9 and 10). We recommend using large-capacity HDD (18TB on our current system) for image acquisition, and cloud platforms for image storage.
6. Click Start Acquisition in the "Acquisition Tab to commence long-term multi-dimensional image capture (Figure 2B).

7. Opening raw Micro-Manager images in CytoSHOW

1. Download the software bundle from http://dispimlineage.wormguides.org.
   NOTE: The software bundle will be downloaded as a .zip file and will need to be extracted to the “C:” directory before use. Further details for installation are given at http://dispimlineage.wormguides.org/diSPIMlineaging_InstallationInstructions.htm .
2. Double-click the file C:CytoSHOWextrasForCCytoSHOW_APP.jnlp to begin running CytoSHOW.
3. Choose menu File > Nouveau > diSPIM Monitor (Micro-Manager). Locate the root data set folder where acquisition timepoint folders were saved. Select any timepoint folder and click Open. Multidimensional navigation windows (called diSPIM monitor windows) are opened automatically for both SPIMA and SPIMB (Figure 3A).
   NOTE: These windows will monitor the root data folder for newly saved raw timepoint stacks (in the event that a specimen is still being recorded). After each new timepoint is acquired, each of the windows monitoring the distinct SPIM arms and sample positions will refresh to display the entire multichannel 4D dataset for each embryo.

8. Generating Max projection images with CytoSHOW

Even before deconvolution, the raw data can be quickly processed to appraise the specimen's global features.

1. Click the Z-MIP button on the left side panel of the image window (Figure 3A, red rectangle) to make maximum intensity projections through the full depth and full-time course of a given position or SPIM­ arm. A Z-projection hyperstack window will appear.
2. In the Z-projection hyperstack window, set projection type to Max intensity. Specify which channels, slices, and time-point frames to process based on user preference.
3. Click OK when completed.
4. Select the folder location to save the Max intensity outputs from the file dialogue window, and then click OK. Allow some time (15–20 minutes, depending on size of dataset and processing power of the computer) for CytoSHOW to generate projection images.

9. Analyzing cell lineages in isotropic high-resolution volumetric data

1. Optional: With the raw data open via diSPIM monitor in CytoSHOW, select the Polygon selection tool (Figure 3A, black arrows) and click just outside the embryo’s anterior, posterior, dorsal and ventral edges (in that exact order) to generate a “bowtie" pattern over the embryo. Do for both views (SPIM-A and SPIM-B, Figure 3A).
   NOTE: This selection specifies the elliptical region of interest (ROI) in which the embryo is centered and records the anterior-posterior axis of the embryo. The bowtie pattern cues CytoSHOW that the user plans to further specify a precise rotation of the final isotropically fused volumes into an orientation that is optimal for lineaging analyses by StarryNite/AceTree. In cases where StarryNite lineaging is not part of the experimental plan, other selection tools and shapes may be chosen to set the ROI for image processing.
2. If multiple embryos were simultaneously imaged by using the multi-position acquisition option, open and perform Step 9.1 for all embryos. This will allow parallel execution of future steps for all embryos in one session. Close the SIMA and SPIMB windows for any embryos you do not wish to process.
3. Click the diSPIM button on the left side panel of diSPIM monitor window (Figure 3A, highlighted in yellow). This reveals a subpanel of controls specific to diSPIM processing.
4. Align green and red channels for each SPIM arm. Because emission channel splitters are used to capture distinct red and green images simultaneously on the same camera, it is important to visually align the exact pixel registration of these two physically adjacent image fields when they are overlaid. Reuse of the same alignment adjustments is typically feasible over many consecutive imaging sessions, but should be checked (as in Steps 9.4.1–9.4.5).
   1. Starting with the SPIMA panel, select the red channel by moving the CM scroll bar to the left (Figure 3A, orange arrow, left panel).
   2. Using the x-, y-, and z-adjusters (Figure 3A, orange square), shift the red channel to match the green.
   3. Click diSPIM button (Figure 3A, highlighted in yellow), to close the subpanel and trigger propagation of the same shifts to all other Poste windows.
   4. Confirm that correct alignment propagated to other frames and time points by moving the “z” scroll bar (Figure 3A, blue arrow, left panel) and/or “t” scroll bar (Figure 3A, green arrow, left panel). If multi-position acquisition was performed and several embryos were imaged (Step 5.3), the alignment should have also propagated to those embryos. Confirm by also examining the numbers for x-, y-, and z-adjusters (Figure 3A, orange square, which should be the same for the SPIMA panel of all embryos).
   5. Repeat Steps 9.4.1–9.4.4 for the SPIMB diSPIM monitor window (Figure 3A, right panel).
5. Click the “diSPIM” button and then the “Fuse” button (Figure 3A, blue rectangle) to open a dialog box called “Deconvolve/Fuse diSPIM Raw Data Volumes” (Figure 3B). Set parameters as shown in Figure 3B. These parameters are briefly addressed in the following substeps:
   1. Set Key registration on Channel 1 (488 nm laser) or 2 (561 nm laser). Select the channel with denser or more ubiquitous signal. For lineaging experiments always select the channel used to image the ubiquitous nuclear histone fluorescence.
   2. Set input volumes relative orientation to +1 or -1. The correct orientation index depends on a specific diSPIM’s camera placements (Figure 4).
      NOTE: If uncertain, test each option by duplicating a single timepoint from both SPIM A and B diSPIM monitor window, following steps 9.1–9.12 and arbitrarily choosing an input volumes relative orientation to test. Incorrect orientations will yield blurry images with artifacts, while correct orientations will yield clear images. The input volumes relative orientation value that yields the clear image can then be reused for all future data from the given diSPIM instrument.
   3. Choose whether the fused volume should be oriented the same as the A or B input volume (based on user preference).
   4. Select "Fresh registration for every volume". This option controls how SpimFusion calculates registration matrices for each volume pair at each timepoint. The “Fresh” option allows the algorithm to optimize registration adaptively at each timepoint.
   5. Set the number of deconvolution iterations to 10. This number tends to reliably yield the desired high resolution in a time-efficient manner.
   6. Optional: If auto-lineaging is desired (highly recommended), check Auto-launch StarryNite lineaging of Fused Volumes. This option will launch StarryNite automatically to segment and track cells in the imaged volumes produced by SpimFusion.
   7. Optional: For highest accuracy in automated lineaging, it is best to reposition the isotropically fused embryo volumes into the canonical "ADL" (Anterior[x-west], Dorsal[y-north], Left[z-near]) orientation. Select the Define volume output orientation in preview option to indicate this choice. CytoSHOW will respond by processing an initial pair of isotropically fused volumes, allowing the user to closely observe and specify the rotations needed to achieve ADL registration.
   8. Click Yes once all parameters are selected.
6. Specify the Output directory in which to save the processed files. Click OK.
7. Optional: If the Define volume output orientation in preview option was selected, set the t scroll bar (Figure 3A, green arrow, left panel) in the SPIM-A window to the early timepoint at which ABa and ABp cells have reached metaphase. Set the t scroll bar in the SPIM-B window to the later comma stage of development. This will aid in specifying ADL orientation.
8. Optional: Click OK when ready. If the previewing option in 9.5.7 above was selected, only two preview volumes will be isotropically fused for the timepoints indicated by the t-sliders of the SPIM-A and SPIM-B image windows. These two preview timepoints can be used to specify precise realignment of output embryo volumes to the ADL orientation, as explained below.
   1. Locate the newly displayed 3DProjY_Decon-Fuse_…. window. Move the t scroll bar to time point 2 of this preview window. Move the Z slider until the view directly down the embryo's long axis is shown.
   2. Move the t scroll bar back to time point 1 of the 3DProjY_Decon-Fuse_…. window. Choose the line-selection tool and draw a line selection from the EMS cell (ventral-most round nucleus) through the plane of the AB-cell metaphase plates.
   3. Click the orange diSPIM Preview button on the 3DProjY_Decon-Fuse_…. window. The fine adjustments to the orientation of the previewed imaged volume will be saved for use in processing the full dataset.
9. Optional: The Deconvolve/Fuse diSPIM Raw Data Volumes dialog will reappear, just as in step 9.5 above. Click Yes without choosing the Define volume output orientation in preview option. Specify the Output Folder for the full data processing run.
10. Set the t scroll bars (Figure 3A, green arrow, left panel) of the diSPIM monitor windows to the starting timepoint (SPIMA) and ending timepoint (SPIMB) of the full span of images to process. Then click Ok.
11. As SpimFusion progresses, CytoSHOW opens and refreshes a multidimensional window showing the sliced-4D isotropic fused volume for each embryo, as well as two windows with rotating-4D max-intensity projections of the isotropic volume. During this time, do not disrupt or close any CytoSHOW window until isotropic fusion and lineage tracking are complete.
12. Optional: Note that once the StarryNite splash screen has appeared and later disappeared, the full data-processing pipeline has been completed. This window must not be closed during processing or StarryNite will be interrupted.

10. Opening StarryNite lineage trace series in AceTree (optional)

1. Open the customized version of “AceTree_16BitCompat.jar” provided.
2. Choose menu the File > Open configure file. Locate your Output directory previously indicated to CytoSHOW. Open the Decon_Fuse_…_Pos[n] subfolder for embryo [n]. Select aaa_edited.xml and Open.
3. Use the AceTree menu Edit > Edit tools to open the Edit Track and Adjust or Delete Cells windows.
4. Click the half-shaded circle Figure 5B, red square to adjust the Red and Green intensities.
5. Proceed with lineage visualization and editing as previously described^5,6,8 (manuscripts are also included in our download bundle).