Imaging and Quantification of Immune Cells in the Tumor Microenvironment Using a Zebrafish Model of Neuroblastoma

Summary

This protocol describes methods for generating compound transgenic zebrafish, imaging immune cells associated with tumors in larval and adult zebrafish, and quantifying immune cell frequency within the tumor microenvironment.

Abstract

The recruitment of immune cells to the tumor microenvironment (TME) is an important component of immune regulation and tumor progression. Studying the spatial and temporal relationships between immune and tumor cells in vivo can provide insight into interactions within the TME. However, visualizing and quantifying these interactions in intact organisms remains challenging using conventional imaging and histological approaches. Zebrafish provide a valuable model for cancer research owing to their genetic tractability, rapid development, and optical transparency during early developmental stages, enabling visualization of fluorescently labeled cells in vivo. Here, we present a protocol to generate, identify, image, and quantify immune and tumor cells that are differentially labeled with fluorescent markers in a zebrafish model of neuroblastoma. The protocol includes breeding and screening of compound transgenic zebrafish, live imaging of larvae, preparation and imaging of adult tumor-bearing fish, and quantification of immune cells surrounding and within tumor regions. Using transgenic reporter lines, CD4⁺ immune cells and MYCN-driven neuroblastoma cells can be visualized and analyzed at different developmental stages. The workflow enables assessment of immune cell localization relative to tumors in both larval and adult fish and can be used to compare immune cell abundance across disease stages. This protocol provides a reproducible approach for imaging and quantifying immune cells within the TME in vivo using zebrafish and may facilitate studies of immune–tumor interactions in other cancer models.

Introduction

The tumor microenvironment (TME) is an ecosystem composed of tumor, immune, fibroblast, and vascular cells¹. The TME plays an essential role in cancer progression, therapeutic response, and anti-tumor immunity. Within the TME, immune cells are key regulators of tumor fate, either participating in anti-tumor responses or inducing immunosuppression². Understanding the recruitment, distribution, and function of immune cells within the TME is critical for elucidating the mechanisms by which tumors evade immune surveillance. However, conventional approaches to studying the TME, such as endpoint histology, flow cytometry, and single-cell sequencing, provide only static snapshots and often require disruption of tissue structure, resulting in the loss of spatial information and cell-cell interactions^3,4,5,6,7. Moreover, these methods are limited in their ability to visualize and quantify immune cells within the TME, especially during the early stages of tumor development.

Zebrafish (Danio rerio) are a powerful model for cancer research due to their low cost, rapid development, relative transparency, and high genetic conservation^8,9. These advantages provide a reproducible and scalable experimental system that allows large sample sizes and efficient microscopy screening within a short time frame¹⁰. Importantly, zebrafish share significant immunological conservation with humans, including key components of both innate and adaptive immune systems, making findings relevant to human disease¹¹. These properties are particularly useful for studying pediatric cancers such as neuroblastoma⁸. Consequently, numerous zebrafish immune-reporter lines and cancer models have been developed¹¹. Therefore, zebrafish serve as a valuable in vivo model for studying interactions between tumor and immune cells, offering insight into how immune regulation shapes tumor development, particularly during early stages.

Previous studies investigating the TME in zebrafish have heavily relied on tumor xenograft models combined with endpoint imaging to assess immune cell localization within tumors^{12,13,14,15,16,17}. While these methods provide useful insights, they are limited by their reliance on two-dimensional imaging, which cannot fully capture the three-dimensional organization of the TME and the spatial relationships between tumor and immune cells during development¹⁴. In addition, tumor xenograft models involve the artificial transplantation of human tumor cells and may not fully recapitulate natural tumor initiation¹⁸. These models are also typically short-term and cannot fully capture tumor progression or changes in the spatial relationship between tumor and immune cells during development^14,19. Furthermore, this approach provides a limited representation of the spatial organization, developmental context, and physiological relevance of immune cell behavior within the TME. Recently, we and others have applied compound transgenic zebrafish, in which tumor and immune cells are differentially labeled with fluorescent markers, to visualize immune cells associated with tumors^20,21. These compound transgenic fish enable the study of the TME throughout animal and tumor development, facilitating investigations of immune-tumor interactions in vivo.

Here, we present a method that utilizes confocal microscopy to study immune cells within the TME of a zebrafish neuroblastoma model. In this approach, we used zebrafish reporter lines that label immune cells and a genetically engineered zebrafish line that overexpresses the human MYCN oncogene under the dopamine-β-hydroxylase promoter (dβh), resulting in the development of neuroblastoma resembling human disease²⁰. In this compound transgenic zebrafish model, tumor and immune cells are differentially labeled with EGFP and mCherry, respectively. The dβh promoter is active in sympathoadrenal lineage cells and has been used in multiple zebrafish models of neuroblastoma^22,23,24,25. Additionally, we utilized a Cd4 promoter driving mCherry fluorescence to visualize CD4⁺ cells¹⁵. Confocal imaging with Z-stack acquisition was applied to visualize the TME in three dimensions at different stages of tumor development, enabling assessment of the spatial relationships between tumor and immune cells within intact tissue. This method enables analysis of the organization of immune cells relative to tumor cells during different stages of tumor development.

Protocol

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at Boston University Chobanian & Avedisian School of Medicine under protocol PROTO202100001. The overall workflow for generation of compound transgenic zebrafish, larval imaging, adult tissue preparation, and image analysis is summarized in Figure 1.

Figure 1. Workflow for visualization and quantification of immune cells within the zebrafish tumor microenvironment (TME). Schematic overview of the experimental workflow. Immune-reporter zebrafish were crossed with tumor-prone zebrafish to generate compound transgenic offspring expressing fluorescent markers in both immune cells (red) and tumor cells (green). Embryos were screened by fluorescence microscopy to identify compound transgenic fish. Selected fish were either subjected to live confocal imaging during larval development (<21 days post-fertilization [dpf]) or processed by fixation, cryosectioning, and confocal imaging for fish older than 21 dpf. This workflow enables visualization and quantification of immune cells associated with the TME across multiple developmental stages. Part of the illustration was created using BioRender. Please click here to view a larger version of this figure.

1. Generation of Compound Transgenic Zebrafish Lines Through Breeding

1. Select adult immune-reporter fish and tumor fish that are at least 3 months old for natural breeding.
   NOTE: For the representative results, Tg(cd4-1:mCherry) heterozygous fish were crossed with Tg(dβh:MYCN; dβh:EGFP) heterozygous fish to differentially label immune cells and tumor cells^15,23.
2. On the evening before breeding, fill a two-chamber mating tank two-thirds full with fish water.
3. Place a grated top insert into the mating chamber. Use dividers to separate one male from one or two females.
4. Leave the dividers in place overnight. Add artificial plants or other enrichment items to simulate natural breeding conditions.
   NOTE: A 1:2 male-to-female ratio increases the likelihood of successful spawning when breeding tumor male fish with immune-reporter fish. Do not breed two males with one female, as competition between males may occur.
5. On the following morning, remove the dividers after the facility lights turn on.
6. Tilt the grated top chamber slightly using the divider as a wedge to create a shallow water area. Allow the fish to spawn for at least 1 h.
7. Collect eggs from the bottom of the mating chamber by pouring the water through a fine-mesh net.
8. Transfer the eggs to a sterile Petri dish containing fish water at a density no greater than 1 egg per 50 mm² (approximately 200 eggs per 100 mm dish).
9. Remove dead and unfertilized eggs.
   NOTE: Healthy fertilized eggs are translucent and round. Remove cloudy, white, unfertilized, or deformed eggs.
10. Double-check the fish to avoid mixing sexes or cross-contaminating transgenic tanks.
11. Return the males and females to their respective tanks.
12. Place the embryos in a 28.5°C incubator overnight. Do not exceed the embryo density specified in Step 1.8.
13. Bleach the embryos on the following morning to remove contaminants.
    1. Prepare a working bleach solution consisting of 100 µL bleach in 200 mL system water. Prepare a neutralization solution consisting of 0.5 g sodium thiosulfate in 150–200 mL system water.
    2. Sort and transfer viable embryos into a mesh-net well insert. Submerge the insert in the bleach solution for 5 min.
    3. Transfer the insert directly into the sodium thiosulfate solution. Incubate for 2 min to neutralize residual bleach.
    4. Rinse the embryos in a final bath of fresh system water. Use a squirt bottle to gently transfer the embryos into clean petri dishes containing fresh system water.
14. Return the embryos to the incubator after bleaching.
15. Transfer larvae at 5–7 days post-fertilization (dpf) or later to a larger tank containing 5 ppt rotifer water for rearing.
    1. Prepare 5 ppt rotifer water by mixing 1 part stock rotifer water (15 ppt) with 2 parts zebrafish system water.
    2. Verify that the final salinity is 5 ppt using a conductivity meter before transferring the larvae.
       NOTE: Sorting EGFP+ MYCN larvae at 3–5 dpf facilitates identification of tumor-prone fish before rearing. Sorting MYCN embryos at this stage avoids false-negative identification because EGFP expression in neural crest cells may decrease later in development due to MYCN-induced apoptosis.

2. Screening Zebrafish for Fluorescent Markers

1. Prepare a 0.4% (w/v) tricaine stock solution using system water.
2. Transfer the fish to a clean petri dish containing only enough system water to cover the bottom.
3. Add the tricaine stock solution to achieve a final concentration of approximately 0.005% (50 mg/L) for fish younger than 14 dpf or 0.012% (120 mg/L) for fish older than 14 dpf. Confirm adequate anesthesia by observing little to no voluntary movement.
4. Remove excess water from the dish once the fish are anesthetized and no longer capable of swimming while maintaining visible opercular or cardiac movement.
5. Screen the fish for fluorescence. Identify green or red fluorescence in the expected region (e.g., the interrenal gland region in MYCN fish).
6. Separate the fish into groups using a stiff-bristled sorting tool.
   NOTE: Sort tumor-prone fish as early as possible because MYCN-induced apoptosis during later developmental stages may result in false-negative identification. Sort immune-reporter fish immediately before imaging at the larval stage or between 2 and 4 months of age for adult studies. This approach minimizes handling-associated stress and maximizes survival.
7. Transfer positive and negative fish to separate tanks containing system water for continued rearing.
   NOTE: Step 3 describes handling and imaging of larvae younger than 21 dpf. Steps 4–6 describe fixation, tissue preparation, imaging, and image analysis of older fish.

3. Larval Live-Mounting and Imaging

1. Place sorted larvae in a 100 mm petri dish containing tricaine fish water (see Step 2.3 for age-appropriate concentrations).
2. Transfer the sedated larvae to a glass-bottom imaging dish.
3. Use only enough solution to cover the larva and maintain sedation.
4. Prepare 1%–1.5% low-melting agarose in fish water. Heat the agarose solution until fully dissolved.
5. Allow the agarose to cool for 1–2 min. Maintain the agarose at 30°C using a heat block to prevent solidification prior to mounting.
6. Position each larva laterally in the imaging dish.
7. Apply approximately 30–200 µL of agarose over the larva to immobilize it. Use larger volumes for larger larvae and smaller volumes for smaller larvae.
8. Once the agarose has solidified, add approximately 100 µL of 0.005% tricaine solution over the agarose.
9. Image the larvae using a confocal microscope.
   1. Use a standard laser-scanning confocal microscope with a resonant scanner. Use a 40× objective with a numerical aperture greater than 1.10.
   2. Set the pinhole to 1.0 Airy unit (AU) for all imaging channels.
   3. Use a 488 nm laser for EGFP excitation and a 594 nm laser for mCherry excitation. Keep laser output below 25 mW to minimize photobleaching and phototoxicity. Adjust detector gain and laser power using a range indicator lookup table (LUT) to avoid pixel saturation and black clipping. Do not exceed a detector gain of 800 V because higher voltages introduce excessive noise.
   4. Use a voxel size of 0.42 × 0.42 × 6.0 µm (x, y, z) at a resolution of 1024 × 1024 pixels and a Z-step interval of 6.0 µm. Acquire optical sections continuously through the full volume of interest, typically spanning 90–120 µm in total imaging depth. Use highly sensitive photodetectors operating in 16-bit acquisition mode.
   5. Acquire images using unidirectional line scanning at a resonant scan frequency of 33.3 kHz (approximately 1.58 µs pixel dwell time). Use 2× or 4× averaging to reduce motion artifacts while limiting laser exposure.
      NOTE: Optimize tricaine exposure to maintain larval survival during imaging. After imaging, add fish water until the imaging dish is approximately half full.
10. Add fresh fish water to the imaging dish immediately after imaging.
11. Gently remove the larva from the agarose using a stiff-bristled stick and transfer the larva to a dish containing clean fish water for recovery.
12. Return the fish to its original tank once normal swimming behavior resumes.
    NOTE: For fish older than 21 dpf, the skin overlying the tumor must be removed by cryosectioning to enable high-resolution confocal imaging.

4. Fish Fixation and Cryosectioning

1. Euthanize fish in accordance with approved institutional animal care guidelines.
2. Make a ventral incision extending from the anal pore to the pectoral girdle to expose the abdominal cavity and facilitate fixative penetration into internal tissues.
3. Transfer the fish using forceps into a 1.5 mL tube containing 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) or a volume of fixative at least 10-fold greater than the tissue volume.
   NOTE: Avoid handling the region containing the tumor to preserve tissue integrity.
4. Place the samples on a nutating rocker set to gentle continuous agitation (30 rpm) and incubate overnight for at least 12 h at 4°C.
   CAUTION: PFA is hazardous, toxic, and potentially carcinogenic. Wear appropriate personal protective equipment, including gloves and eye protection. Perform all procedures involving PFA in a fume hood and avoid inhalation of vapors.
5. Remove the PFA and wash the samples at least four times with 1.5 mL of 1× PBS. Gently invert the tube two to three times during each wash.
6. Transfer the fish to 30% sucrose prepared in PBS.
7. Incubate the fish in a cold room overnight until the specimen sinks to the bottom of an upright tube (approximately 12 h), indicating adequate sucrose penetration.
8. Remove excess sucrose solution by gently blotting the fish on a laboratory tissue.
9. Transfer the fish to storage tubes.
10. Store the samples at −80°C until imaging.
    NOTE: Process samples within 3 months to minimize fluorescent protein degradation and signal loss.
11. Transport the fixed fish to the cryostat room on dry ice.
12. Place the fish flat in a plastic mold with one sagittal side facing downward.
13. Add Optimal Cutting Temperature (OCT) compound until the fish is fully covered.
14. Freeze the mold at −20°C or in the cryostat until the OCT turns white.
    NOTE: Position the fish as flat as possible before freezing to facilitate consistent sectioning.
15. Cryosection the zebrafish in the sagittal plane to remove the skin for subsequent whole-mount imaging.
    1. Cool the cryostat chuck.
    2. Apply room-temperature OCT to the chuck.
    3. Press the OCT fish block firmly into the OCT layer.
    4. Freeze the chuck and block until the newly applied OCT turns white.
    5. Mount the chuck in the cryostat.
    6. Cryosection the block at a thickness of 14 µm until the blade reaches the specimen.
       NOTE: Specimen shifting may occur during freezing, causing the fish to sit higher in the block than expected. Entry into the specimen is indicated by the appearance of translucent tissue replacing the opaque OCT background.
    7. Reduce the section thickness to 7 µm.
    8. Continue sectioning for one additional cut.
       NOTE: This procedure removes the skin while preserving a uniform sagittal profile across the specimen.
    9. Confirm the endpoint by identifying skin in the collected sections.
       ​NOTE: Skin appearance in sections provides a more reliable indicator than eye morphology. Perform this procedure only when the skin obscures imaging of the tumor microenvironment.
16. Warm the chuck gently with your hands.
17. Remove the OCT block from the chuck using forceps. Avoid disturbing the cut surface of the specimen.
18. Trim excess OCT surrounding the fish while the OCT remains frozen.
19. Remove the fish from the OCT block using forceps once the OCT softens to a viscous consistency.

5. Mounting and Imaging of Fish with Skin Removed

1. Place the fish in a glass-bottom dish with the sectioned sagittal side facing the glass surface.
   NOTE: Maintain the orientation of the fish during handling. Identifying the sectioned side may become difficult after the fish is removed from the OCT block.
2. Prepare 1%–1.5% low-melting agarose.
3. Apply approximately 30–50 µL of agarose to the dorsal and ventral sides of the fish to secure the sample.
   NOTE: Avoid placing agarose beneath the fish because it can reduce image quality. Use only the minimum amount of agarose required for stabilization.
4. Keep the sample on ice during transport. Image the sample within 2 h of mounting.
5. Transport the sample to the confocal microscopy facility.
6. Image the sample using confocal microscopy.
   1. Use a standard laser-scanning confocal microscope with a resonant scanner. A two-photon microscope may be used when autofluorescence limits image quality. Use a corrected 20× objective lens with a numerical aperture of at least 0.80.
   2. Set the pinhole to 1.0 Airy unit (AU) for all imaging channels.
   3. Use a 488 nm laser for EGFP excitation and a 594 nm laser for mCherry excitation. Keep laser output below 25 mW. Adjust detector gain and laser power using a range indicator lookup table (LUT) to avoid pixel saturation and black clipping. Do not exceed a detector gain of 800 V because higher voltages introduce excessive noise.
   4. Configure image acquisition parameters as follows.
      1. Use a voxel size of 0.42 × 0.42 × 6.0 µm (x, y, z) at a resolution of 1024 × 1024 pixels and a Z-step interval of 6.0 µm.
      2. Acquire spectral image stacks through the full volume of interest, typically spanning a total Z-stack acquisition range of 90–120 µm (15–20 optical sections).
      3. Initiate image acquisition in the fluid space immediately above the tissue surface and continue through the full target imaging depth to ensure complete coverage of the specimen.
      4. Acquire images using highly sensitive photodetectors operating in 16-bit acquisition mode.
   5. Acquire images using unidirectional line scanning at a resonant scan frequency of 33.3 kHz (approximately 1.58 µs pixel dwell time) with 8× line averaging.

6. Image Analysis and Quantification

1. Image visualization using Fiji (version 1.54p).
   1. Open the confocal z-stack images.
   2. Merge colors to see channel overlay by selecting Image > Color > Merge Channels.
   3. Generate a projection of the full z-range containing the imaging region by selecting Image > Stacks > Z Project. Enter the slice range of interest, as well as Max Intensity projection type. This will display the 2D projection of the image.
      NOTE: For larval fish, it is feasible to capture the whole TME by confocal microscopy. Therefore, we recommend the volumetric normalization and quantification method for the TME analysis. For older fish, it is impossible to capture the whole TME by confocal microscopy; therefore, we recommend applying the area-based normalization and quantification method. Based on our analysis and quantification of the larval fish, both methods can provide reliable biological results.
2. Volumetric normalization and quantification of the TME in the larval fish.
   1. Quantifying tumor volume using MATLAB R2022b (download the package from the Matlab website) and Bio-Formats package (download from the openmicroscopy website).
   2. The user runs a script that performs the following tasks:
      1. Prompt the user to select a .CZI file from their computer.
      2. Read the image and extract the data from the channel containing the tumor region.
      3. Identify the region for each slice that is the tumor region through a threshold of fluorescence (can be through user input or machine learning), keeping the same threshold for all analyzed images for consistency.
      4. Outline the tumor region and display the image with the tumor region outlined in a collage of all slices for user verification.
      5. Calculate the area of the tumor region per slice in pixels and convert to mm² using the scale of the image and imaging settings.
      6. Multiply the area of the tumor region by the depth (stored in the file’s metadata) to get the volume of each slice.
      7. Sum the calculated values of each slice to obtain the total volume of the imaged tumor and its surrounding region in voxels or mm³.
      8. Scale both volume measurements by multiplying by 10⁴ for tumor volume and 10⁶ for the surrounding region. This will create a more reasonable cell number:volume ratio during normalization.
   3. Quantifying cell number.
      1. Open the confocal z-stack images.
      2. Merge colors to see channel overlay by selecting Image > Color > Merge Channels.
      3. Generate a 3D view of the full z-range containing the imaging region by selecting Image > Stacks > 3D Project. Select Brightest Point as the projection method and other settings to the desired values.
      4. Due to the small number of cells, manually count and tabulate the cells, separating cells located within the tumor boundary from those located outside the tumor boundary.
         NOTE: Cells on the boundary of the tumor region are counted as being outside.
   4. Normalization.
      1. For infiltrating cells inside the tumor region, divide the number of cells by the scaled tumor volume. For the cells outside the tumor region, divide the number of cells outside the tumor volume by the scaled volume of the full image minus the tumor volume.
      2. Generate a table that includes the following headings: “fish identification”, “tumor volume”, “# of cells inside tumor region”, “# of cells outside tumor region”, “normalized cells inside”, “normalized cells outside”.
3. Area-based normalization and quantification of the TME.
   1. Define the tumor region of interest (ROI) using the Rectangle or Freehand selection tool to outline the tumor.
   2. Add the ROI to the ROI Manager by pressing T or by selecting Analyze > Tools > ROI Manager > Add.
   3. Measure the ROI area using Analyze > Measure.
   4. Confirm that Area is selected under Set Measurements before recording measurements.
   5. Save the ROI in the ROI Manager.
   6. Apply the same ROI to all images using ROI Manager > More > Multi-Measure.
   7. Do not resize the ROI between images.
   8. Manually count cells within the ROI. If cells are more, count cells using the Cell Counter plugin by selecting Plugins > Analyze > Cell Counter.
   9. Mark each counted cell by left-clicking when its centroid falls within the ROI.
   10. Scroll through the z-stack to verify cell identity.
   11. Avoid counting the same cell more than once across multiple image planes.
   12. Count infiltrating cells separately if they contact or lie within the tumor boundary.
   13. Record infiltrating cells as a distinct category even when the cell centroid falls outside the ROI.
   14. Convert the ROI area from pixels to mm².
   15. Obtain the pixel width, pixel height, and Z-step interval from Image > Properties.
   16. Calculation of normalized cell counts.
       1. Divide the raw cell count for each fish by the corresponding ROI area value.
       2. Record the resulting value as the normalized cell count.
       3. For absolute density measurements, calculate the physical area using the image calibration values obtained from Image > Properties.
       4. Divide the raw cell count by the calculated area (mm²).
       5. Record the result as cell density expressed as cells/mm².

Results

The overall experimental workflow is shown in Figure 1. Adult Tg(cd4-1:mCherry) zebrafish were crossed with Tg(dβh:MYCN;dβh:EGFP) zebrafish to generate compound transgenic offspring in which CD4⁺ cells and dβh-expressing cells were differentially labeled with mCherry and EGFP fluorescence, respectively. Embryos were screened for the appropriate fluorescent markers and subsequently analyzed by either live confocal imaging during larval developmen...

Discussion

The protocol described here utilizes zebrafish as an in vivo model to visualize and quantify immune cells within the TME. Several factors are critical for successful implementation of this approach. First, appropriate selection and validation of compound transgenic zebrafish lines are essential. Larvae should be screened for the expected fluorescent expression patterns and, if required, confirmed by genotyping. Immune-reporter and oncogene expression should be verified prior to experimentation to ensure accurate...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
200 µL Gel Loading Tip	MilliporeSigma	1022-0600	Used for larval manipulation
Artificial Plants	The Hidden Reef	62874200000000000	Optional breeding enrichment
Bleach	Fisher Scientific	50371500	Used for embryo decontamination
Cell Counter Plugin	National Institutes of Health (NIH)	N/A	Fiji plugin used for cell quantification
Computer and Monitor	HP	2TB68A8#ABA	Used for image acquisition and analysis
Confocal Microscope	Zeiss	LSM 710	Used for larval and adult imaging
Cryomold, Plastic	Epredia	22-050-160	Used for OCT embedding
Cryostat	Thermo Fisher Scientific	95-664-0EC70	Used for cryosectioning
Dry Ice	Fisher Scientific	N/A	Used for sample transport
Fiji Software	National Institutes of Health (NIH)	Version 1.54p	Image analysis software
Fine-Mesh Net	Carolina Biological	651345	Used for embryo collection
Fish Incubator	VWR	35960-056	Maintained at 28.5°C
Forceps	Fisher Scientific	10-316B	Used for sample handling
Glass-Bottom Dish	IBIDI	81218-200	Used for imaging
Low-Melting Agarose	Prolab Supply	9012-36-6	Used for larval and adult mounting
Mating Tank (main tank, divider, grated section)	Carolina Biological	161937	Used for zebrafish breeding
Nutating Rocker	Midsci	R2D-30	Used for fixation during overnight incubation
OCT Compound	Tissue-Tek	4583	Embedding medium for cryosectioning
Paraformaldehyde (PFA)	Sigma Aldrich	158127-500G	Fixative
Petri Dish, Plastic	MilliporeSigma	BR452000	Used for embryo and larval handling
Phosphate-Buffered Saline (PBS)	Gibco	14080-055	Used for washing and fixation procedures
Plastic Transfer Pipette	Fisher Scientific	50-998-100	Used for embryo and larval transfer
Razor Blade	Fisher Scientific	12-640	Used during sample preparation
Conductivity Meter	YSI	EC300ACC-04	Used to verify rotifer water salinity
Rotifers	The Hidden Reef	116151603	Used for larval rearing
Sodium Thiosulfate	Thermo Fisher Scientific	AAA1762936	Used for bleach neutralization
Sucrose	Fisher Scientific	S25590A	Cryoprotection reagent
Tricaine Methanesulfonate	Western Chemical	NC0872873	Anesthetic; sold by Fisher Scientific
Tweezers	Fisher Scientific	12-000-122	Used for sample handling
Upright Fluorescence Microscope	Leica	M165	Used for fluorescence screening