Spatially Compact Arrangement of Larval Zebrafish Sections for Spatial Transcriptomic Analysis

Here, we present a method for aligning and cryosectioning multiple Zebrafish (Danio rerio) larvae samples and collecting them on a single slide for spatial transcriptomic analysis.

Spatial transcriptomic techniques are a sophisticated tool in biomedical research to visualize spatially registered gene expression patterns. Imaging and analysis of multiple samples with spatial imaging platforms can be costly. Performing these tests over multiple experimental conditions, as seen in developmental studies, further increases costs. To reduce costs, this study sought to optimize the techniques and strategies of spatial transcriptomic specimen arrangement for developmental studies. Here, the study utilized zebrafish, which are a well-established developmental vertebrate model that is transparent during development, have ~70% genetic homology to humans, and a highly annotated genome ideal for transcriptomic analysis. Because of their small size, developing zebrafish also allows for compact placement of serial sections across several biological replicates. Herein, we report optimized fixation, cryosectioning, and reliable alignment of multiple fish samples within the imaging area of a multiplex in situ hybridization spatial imaging platform. With this method, zebrafish as young as 15 days post fertilization (dpf) from at least 4 different molds and up to 174 sections can be successfully cryosectioned, collected within the imaging area of 22 mm 10.5 mm (for an in situ spatial transcriptomic slide), and processed simultaneously. Based on section quality, sample alignment, and sample size per slide, this method in zebrafish optimizes output and per sample cost of spatial transcriptomic techniques.

Assessment of spatially distinct expression patterns in tissue remains critical for our understanding of genomic influences in development, cancer, and disease^1,2,3. Spatial transcriptomics combines multiplexed expression techniques with the spatial registration of expression in tissues. "Spatial transcriptomics" was first coined by Ståhl and colleagues⁴, where mounted cancer specimens were probed using in situ next-generation sequencing. Since that time, "spatial transcriptomics" has been used as a catch-all for high throughput expression studies combined with spatial registration. While these are powerful tools, they are also expensive undertakings that often require large institutional investment and laboratory costs before data can be generated⁵. Strategies to minimize cost while preserving high-quality data are in high demand.

Zebrafish, Danio rerio, have become an important model system for studying developmental biology and offer a means of multiplying vertebrate whole organ (and organism) analyses in limited space. Zebrafish are small (4-6 mm as larvae and 2-3 cm as adults) and can lay hundreds of transparent eggs at a time⁶. Zebrafish embryos are fertilized externally and develop rapidly, allowing researchers to introduce transgenes at early stages of development to readily generate gain- and loss-of-function alleles⁷. Fitting multiple specimens on a single slide is an appealing strategy to reduce costs. Their high fecundity and small size make zebrafish an ideal candidate for multiplexing spatial transcriptomic assays which have restricted space for specimens⁸.

Cryosectioning zebrafish larvae is a challenging technique. Many spatial transcriptomic platforms have not been optimized for zebrafish paraffin sections and require cryosections when working with zebrafish as a model organism to preserve tissue structure and retain RNA transcripts. Additionally, the small size of zebrafish makes it difficult to obtain quality cryosections and analyze multiple samples effectively. This task becomes more difficult when working with zebrafish larvae that are smaller and more fragile than their adult counterparts. To overcome these challenges, we describe a method that reliably aligns multiple samples and utilizes the imaging area of spatial imaging platforms efficiently to obtain many high-quality sections on a single slide that can then be imaged and analyzed by spatial imaging platforms (Figure 1). In this instance, this method is applied to a spatial transcriptomic imaging platform.

This protocol follows the guidelines of Dartmouth College's institutional animal care and use committee.

1. Preparing cryostat

1. Cool the cryostat to -22 °C and clean the interior surfaces of the cryostat by brushing debris into the receptacle. Place all necessary brushes and tools inside the chamber.

2. Preparing the disposable base mold

1. Prepare a base mold (37 mm 24 mm 5 mm disposable plastic mold) for sample alignment by drawing a straight line across the inside of a base mold with a permanent marker to use as a reference point for sample alignment. Place a piece of lab tape on the inside of the base mold where the straight line should be before drawing a line for best results (Figure 2A).
2. Draw a dot on the inside of the base mold near the left or right wall to ensure proper sample orientation during cryosectioning (Figure 2B).
3. Measure the desired cutting angle with a protractor and mark this on the inside of the base mold for each sample (Figure 2C).
4. Apply a shallow layer of freezing (optical coherence tomography [OCT]) medium to the prepared base mold. Ensure there is just enough freezing medium to cover the samples.
   1. Avoid air bubbles when applying the freezing medium by priming the nozzle of the medium bottle and adding the necessary amount of medium to one corner of the base mold before shifting the horizontal plane of the base mold so that the medium is distributed evenly across the entire surface.
   2. Squeeze the bottle slowly when dispensing the freezing medium.
5. Add ice to a 1 L beaker, place the base mold with freezing medium in the ice bath, and incubate for at least 10 min to cool the medium.

3. Preparing dry ice:100% ethanol bath

1. Prepare a dry ice and 100% ethanol bath in a fume hood by adding one part of 100% ethanol to one part of dry ice in an ice bucket.
2. Use a disposable aluminum dish or fold aluminum foil into a boat large enough to fit a disposable base mold. Ensure that the dish or boat is large enough for the base mold to lay completely flat.
3. Place the dish or boat in the bath and cover the bucket. Allow 5-10 min for the bucket to cool before freezing samples.

4. Euthanizing samples

1. Randomly select zebrafish for sectioning. If samples vary in size, separate them into groups by relative size to make accurate alignment easier (see discussion for details). Place larger fish and smaller fish in separate dishes.
2. Fill a beaker with fish system water and place the beaker in an ice bucket. Surround the beaker with ice.
3. Monitor the temperature of the water with a thermometer. Let the temperature stabilize between 2-4 °C.
4. Use a net or strainer to put one group of fish into the 4 °C water. Fish should be fully immersed in water and not in contact with ice. Once opercular movement has ceased, add ice to the water to ensure it remains below 4 °C. Leave fish in 4 °C water for 10 min.
   NOTE: Continue with the embedding, alignment, and flash freezing of each group of euthanized samples before euthanizing the next group. Replace the water each time.

5. Embedding and alignment

1. Collect the euthanized fish after they have been submerged in 4 °C water for 10 min. Remove the fish from the water with fine-tipped forceps by grabbing them by the caudal fin and dry them by gently pressing them against an absorbent, lint-free wipe.
   NOTE: For high-quality sections, it is critical to limit the time between removing fish from the 4 °C water and flash-freezing
2. Working with the prepared base mold in an ice bath under a stereomicroscope (10x magnification), place each sample into the base mold along the reference points in the correct orientation and gently cover the samples with another thin layer of freezing medium.
   NOTE: Do not fill the whole mold with freezing medium. Instead, only use a thin layer, just enough to cover all the samples.
3. Use an anatomical reference point to precisely align the fish to the lines marked on the inside of the base mold. Use fine-tipped forceps to adjust the orientation of each fish so that they are aligned and in the same orientation. Avoid creating bubbles in the freezing medium by moving slowly.
4. Apply a piece of dry ice on the bottom of the base mold under the samples until they are locally frozen into position. Keep the horizontal plane of the base mold level to avoid shifting the samples off of their reference points before freezing into position with dry ice.
5. Place the base mold with the samples onto the aluminum boat or dish in the dry ice:100% ethanol bath. Ensure the boat is floating at the surface of the bath and the base mold remains dry. Cover the bath and allow samples to float for 10 min.
6. Wrap the frozen base mold in foil and store in -80 °C freezer until ready to section. Repeat steps 4.2-5.6 for any remaining groups.

6. Cryosectioning

1. Bring frozen base molds to the pre-cooled cryostat for cryosectioning and place them in the cryostat chamber. Transport frozen blocks in a box with dry ice to prevent thawing.
2. Remove in situ spatial imaging slide from storage and place it in a pre-chilled slide holder. Store the slide holder with spatial imaging slide in the -22 °C cryostat chamber until ready to collect sections from the region of interest.
3. Remove the frozen samples from the base mold and then freeze them in a chuck with a fresh freezing medium. Freeze them onto the chuck so that the cutting surface will face the blade.
4. Place a fresh, fine microtome blade in the cryostat.
5. Align the mold to the blade and trim the region of interest (recommended trim thickness 20-50 µm). Ensure the markings made inside the mold are transferred to the frozen sample block to help identify the location within the samples.
6. Adjust the mold during the trimming phase so that the cutting surface is parallel to the reference markings in the freezing medium.
7. Collect sections onto a standard positively charged microscope slide and check them by brightfield to confirm when trimming is no longer necessary.
8. Remove the spatial imaging slide from the cryostat chamber and place it in a 4 °C ice bath. Keep the slide in the slide holder. Ensure that the slide does not get wet.
9. Start cryosectioning (recommended 10-14 µm; Figure 3) and collect sections onto positively charged slides until the region of interest in the samples is reached. Check sections by brightfield to confirm the region of interest will be the next section from the mold.
10. Bring the single-cell spatial imaging slide back to the cryostat chamber. Remove the slide from the slide holder and collect sections from the exact area of interest onto the spatial imaging slide row by row using a fine-tipped paintbrush to keep sections from rolling up.
    1. Press a corner of the empty, freezing medium into the knife stage with the backside of the paintbrush so the section remains flat when grabbing the slide for collection.
    2. Use the top of the slide as a pivot point, slowly lower the slide onto the section, and allow sections to adhere to the slide for 3 s before lifting the slide off the knife stage.
    3. Work from left to right when collecting sections in the imaging area of the slide and overlap layers of empty, freezing medium when possible.
    4. Use colored paper borders as a reference to place sections within the imaging area of the slide if tissue is hard to see.
11. After collecting sections from the region of interest, place the slide back into the slide holder. If there are no more samples to be collected onto the spatial imaging slide, store the slide at -80 °C for up to 2 weeks until it is ready for instrument analysis with the in situ spatial imaging platform. If sections need to be collected from multiple molds, put the spatial imaging slide back in the 4 °C ice bath and repeat steps 6.3-6.11 with the next mold.
12. Collect sections before and after region of interest sections from each mold on a standard positively charged microscope slide for hematoxylin and eosin (HE) staining to check that sample alignment and section quality are sufficient before proceeding with analysis.

7. Fixing the sample

1. Remove the reference slides from the cryostat and air dry at RT for 30 min to adhere sections to the slide.
2. Fix sections by placing them in a slide container with 4% paraformaldehyde (Table 1) for 20 min.
3. Wash sections by placing them in a slide container with distilled water for 3 min.
4. Continue with HE staining or dry and store slides at -80 °C for future staining.

8. HE staining of the sections

1. Dehydrate and clean the sections by incubating the slides in 100% ethanol for 2 min, 95% (Table 2) ethanol for 2 min, and then tap water for 1 min. Use a microscope slide staining rack to transfer slides from bath to bath.
2. Stain the nuclei and differentiate by incubating the slides in Hematoxylin for 2 min 45 s, tap water for 1 min, 0.3% acidified alcohol (Table 3) for 1 min, and then running tap water for 1 min. Use a microscope slide staining rack to transfer slides from bath to bath.
3. Stain the cytoplasmic components and dehydrate by incubating the slides in Eosin Y 1% for 45 s, 50% ethanol (Table 4) for 1 min, 95% ethanol for 1 min, and 100% ethanol for 1 min. Use a microscope slide staining rack to transfer slides from bath to bath.
4. Clear the sections by incubating slides in Xylene for 1 min. Mount and cover the slides by applying a drop of mounting medium to the top third of the slide with a transfer pipet and slowly lowering a coverslip on top of the mounting medium with forceps.

9. Spatial transcriptomic imaging and analysis of the sections

1. Remove the imaging slides from -80 °C storage and image with an in situ spatial imaging platform for spatial transcriptomic analysis.
   NOTE: The exact steps involved in imaging will be determined by the spatial imaging platform.
2. Review the quality control metrics of the spatial transcriptomic platform. Important metrics to check are the number of cells detected, median transcripts per cell, nuclear transcripts per 100 µm², and total high-quality decoded transcripts of each gene in the probe set.
   NOTE: These quality control metrics do not have universal thresholds, and expectations for these thresholds will vary depending on the sample and gene panel being used.
3. Read the data output of detectable RNA transcripts, determine which transcripts are low quality based on the experiment's quality control metrics, and filter out low-quality transcripts. Analyze the remaining high-quality transcripts in relation to their spatial arrangement within the section.
4. View the cellular segmentation of the sections and cluster cells based on experimental interests. Compare the RNA transcripts within cell clusters of the region of interest across groups of different ages of zebrafish on the same slide.

In this method (Figure 1), zebrafish is used as an animal model to probe for spatially resolved gene expression patterns. Cryosectioning larval zebrafish efficiently for spatial imaging is challenging. Sections must be high quality to retain tissue structure and detectable genes (Figure 4). Sections containing multiple samples for spatially efficient imaging must be aligned precisely to analyze regions of interest across all samples (Figure ...

This report provides detailed solutions to many of the technical challenges associated with zebrafish as a model organism in spatial transcriptomic analysis during development. In addressing these challenges, our compact specimen arrangement optimizes costs on the emerging spatial transcriptomic platforms¹. Cryosectioning larval zebrafish for spatial imaging is challenging. Sections should retain sufficient tissue structure and transcript quality for satisfactory experimental execution and downstr...

The authors have no disclosures or conflicts of interest regarding this report.

Sectioning and imaging were performed with instruments provided by shared resources at the Dartmouth Cancer Center, funded by NCI Cancer Center Support Grant 5P30CA023108, and the Center for Quantitative Biology at Dartmouth College (NIGMS COBRE).