En face Cryosectioning of Mouse Retina for High-dimensional Spatial Molecular Analysis

Summary

The retina exhibits complex spatial organization across its laminar structure. Modern spatial transcriptomics and sequencing techniques require tissue sections < 20 µm thick, limiting their retinal research applicability. This fresh-frozen cryosectioning method produces thin en face sections of mouse retina while preserving planar spatial relationships, enabling comprehensive molecular mapping across laminae.

Abstract

The complex laminar structure of the retina presents significant challenges for comprehensive spatial molecular mapping. Current techniques for investigating tissue architecture and molecular interactions are limited by technical constraints that compromise spatial relationships or restrict molecular profiling. Existing methods like cross-sectioning, whole mount preparations, chemical or mechanical dissociations, and thick en face sections either disrupt tissue integrity, lose critical spatial context, or most often, are simply incompatible with high-dimensional spatial sequencing platforms.

This study introduces a cryosectioning technique that produces thin (<20 µm) en face sections while preserving the spatial relationships essential for mapping retinal cell distributions. The method maintains RNA integrity and tissue architecture while generating sections compatible with high-throughput molecular analysis platforms. RNA integrity is maintained because this technique sections fresh frozen tissue, which manufacturers of spatial transcriptomic platforms officially support relative to postfixed tissue. We validate this approach using multiple molecular profiling methods, demonstrating successful integration with both manual RNAscope in situ hybridization (12 targets) and Xenium spatial sequencing (300 targets). The technique consistently preserves tissue architecture across samples while maintaining RNA quality suitable for these sensitive molecular applications.

This methodological advancement enables new investigations into retinal cell type distributions, molecular gradients across laminae, and spatial aspects of retinal pathology. The technique's versatility and compatibility with modern spatial biology platforms provide a foundation for comprehensive molecular mapping of the retina's complex cellular organization.

Introduction

Retinal neurons have evolved spatial topographies that facilitate information processing, which in many species, results in gross specializations like foveas, area centralis, and streaks^1,2,3. In contrast to the well-understood diversity of photoreceptor distribution, spatial specialization of retinal output, formed by Retinal Ganglion Cells (RGCs), is less clear. For most species, the RGCs' spatial distributions are entirely unknown. Even for the mouse, arguably one if not the best-studied vertebrate model of visual function, less than a third of RGC subtypes have been mapped with traditional techniques like whole mount immunohistochemistry². These techniques fail because most RGC subtypes do not have a specific genetic marker. Instead, higher-dimensional molecular information is needed for accurate RGC classification⁴ via single-cell RNA sequencing^5,6,7.

Information about RGC distribution is important, as the topographies observed in the mouse demonstrate clear ecological correlates between RGC placement and visual function. The most striking example is the spatial organization of the ɑON-S RGCs, which cluster in the temporal retina. This region receives the image of the prey⁸ during hunting behaviors⁹. It is also well appreciated that RGC subtypes are differentially susceptible to disease¹⁰, and some common diseases like glaucoma have a spatial component to their progression^11,12. Thus, an important question facing the field is: what is the spatial organization of RGC subtypes as they tile the retina¹? This gap in knowledge is currently limiting insights into other visually guided behaviors and diseases.

The focus of the present method is to address these molecular knowledge gaps through an en face cryosectioning approach that preserves spatial relationships while enabling high-resolution molecular analysis. Modern spatial sequencing platforms like 10X's Xenium represent a transformative advancement in molecular profiling, offering unprecedented insights into tissue organization and cellular interactions. However, these platforms, along with lower-plex techniques like RNAscope and spatially barcoded sequencing methods like Visium, share a critical technical constraint: they require tissue sections thinner than 20 µm. This requirement presents a particular challenge for retinal tissue, which typically measures 200 µm in thickness.

To date, researchers have primarily relied on cross sections when applying these technologies to retinal tissue¹³. While cross sections effectively reveal molecular relationships within nuclear and plexiform layers in the context of vertical circuits, they are less suited to capturing the broader spatial distributions of retinal cells that whole mount preparations have traditionally revealed⁸. Recent attempts at en face sectioning have been limited by section thickness constraints, with studies achieving only 50 µm sections¹⁴-too thick for modern high-plex molecular analysis platforms. Other historical approaches that have been deployed to study individual retinal lamina but are not compatible with modern spatial sequencing techniques include enzymatic digestion, combined with mechanical dissociation^15,16, or mechanical dissociations alone^17,18,19,20. Here, we present a novel cryosectioning technique specifically designed for thin, curved retinal tissue that overcomes these limitations. Briefly, our method involves extracting the posterior chamber of the eye, creating relief cuts through both the retina and sclera, flattening the tissue with inner retina against a coverglass, embedding the tissue in OCT, and finally, cryosectioning from outer to inner retina (Figure 1). Our method maintains spatial relationships while producing sections thin enough (<20 µm) for high-dimensional spatial biology tools, as validated through successful implementation with both manual RNAscope (12 targets) and the automated Xenium platform (300 targets), demonstrating compatibility across different complexity scales of molecular analysis.

Protocol

All experiments were conducted in accordance and with approval from the Institutional Animal Care and Use Committee (IACUC) and relevant institutional guidelines approving these methods overseen by the University of Colorado. All steps related to animal use and euthanasia must follow local regulations and experimental permits. All mice were housed with standard care in an accredited veterinary facility with free access to food and water. This en face cryosectioning technique builds on standard retinal dissection techniques optimized for C57BL/6 mice. To develop this technique approximately 150 mice of both sex ranging in age from 6 weeks to 2 years of age were used. The final technique communicated here has a success rate of approximately 95% regardless of mouse demographic characteristics. Note that if a temperature is not listed for a step, room temperature of approximately 20-25 °C is anticipated. Solutions without temperature listed will function if chilled and thus the working temperature range for solutions is 4-25 °C.

1. Dissection

1. Anesthetize the mouse with isoflurane and validate anesthesia with toe pinch, or follow local rules and regulations to induce and confirm anesthesia as other methods are equally appropriate.
2. Euthanize the mouse by cervical dislocation: secure the mouse's head with one hand and the base of the tail with the other, then pull in opposite directions. Perform secondary euthanasia by decapitation.
3. Brand the temporal pole of the globe on the cornea to maintain orientation and create a weak point for dissection (Figure 2B and Figure 3A).
4. Submerge the globe in Ames solution in a dish containing a piece of filter paper (larger than the globe but smaller than the dish).
   NOTE: Use of solutions other than Ames will result in retinal deterioration (Figure 4A).
5. Using sharp #5 forceps and iris scissors, carefully remove all musculature, fascia, vasculature, and nerves from the globe (Figure 2E,F and Figure 3B) without puncturing the sclera (Figure 2D).
   NOTE: Puncturing the sclera during this step (Figure 4B) will result in tissue herniation (Figure 4C).
6. Position the tip of one iris scissor blade perpendicular to the corneal brand, with scissors parallel to the optic nerve head. Puncture the globe at this position (Figure 3C,D).
7. Make a single clean incision through the cornea and limbus, continuing into the sclera and retina toward the optic nerve head (Figure 3E).
   NOTE: Do not cut through the optic nerve head or the sclera and retina will separate. The exact length of this incision will vary depending on the size of the globe, which can vary dramatically between old vs young animals, as well as in some Cre lines relative to wild type animals. We recommend leaving roughly 1 mm of tissue between the deepest extent of this incision and the optic nerve head.
8. Position the blade perpendicular to the initial cut (Figure 3F,G), approximately 1 mm posterior to the limbus (Figure 2C). Begin cutting radially through the sclera and retina.
   1. Maintain a consistent cutting line posterior and parallel to the limbus by using sharp forceps to guide and rotate the globe. While minimizing cuts is ideal, prioritize maintaining a clean radial path. If retina and sclera separate, reposition the iris scissor blade to cut them together.
      NOTE: Cutting into the limbus may cause retinal separation from the sclera during lens extraction. Small snips and rotations are acceptable. Avoid applying force to secure it, as this risks retinal herniation.
   2. Estimate the 1 mm posterior distance using the middle width of the iris scissor blade. When uncertain, err toward a more posterior position, as the retina's connection to the limbus curves internally. A more posterior cut facilitates flat retinal extraction.
9. After completing the radial cut parallel to the limbus, inspect for remaining retinal connections. Carefully sever these with iris scissors until the anterior and posterior globe sections are separate.
10. Using two pairs of sharp #5 forceps, gently separate the anterior and posterior portions of the globe (Figure 3H). In older mice, transparent fibers may connect the lens to the retinal cup; carefully sever these with iris scissors while minimizing retinal-scleral separation.
    NOTE: The lens should remain attached to the limbus and cornea.
11. Using the filter paper as a support platform, cut the retina into a clover leaf (also known as iron cross) pattern with iris scissors (Figure 3I-K).
    NOTE: Clover leaf incisions should be shorter than the primary orienting incision initially made through the limbus if recording orientation with this slice is desired. We recommend reducing the size of these three new radial incisions to be roughly 1-2 mm shorter than the primary incision. If a different feature, like the opsin gradient, will be used for orientation, it is appropriate for all incisions to be the same length while always retaining approximately 1 mm of tissue between the deepest part of the incision and optic nerve head to prevent separation fo the retina and sclera.
12. Repeat the dissection with the second eye. To minimize necrosis, complete anterior chamber removal of the second eye within 10 min of euthanasia.

2. Cryopreservation

1. Prepare a disposable transfer plastic pipette by cutting the tip to create an opening larger than the dissected retina and sclera (hereafter referred to as the retina).
2. Gently suction the retina into the pipette and deposit it onto a glass microscope slide. Using a laboratory wipe, carefully remove excess Ames solution around the retina until it can no longer float freely and can be flattened.
3. Using two sharp #5 forceps, unfold the retina and position it flat with the inner eye cup facing upward. Depending on the size of the tissue, make 0.5-2 mm relief cuts in each cloverleaf segment to further flatten the retina, carefully adjusting the tissue to prevent folding (Figure 5A). Remove additional Ames solution until liquid remains only adjacent to the retina.
4. Using a coverslip, quickly press and transfer the retina (Figure 5B) with its surrounding liquid so that the outer sclera faces upward and the inner eye cup contacts the coverslip, that is, the retinal ganglion cells will face up (Figure 5C).
5. Verify the retina is not folded and make adjustments as needed; add Ames solution for easier manipulation. As excess liquid will cause the tissue to curl, use a laboratory wipe to remove all excess Ames solution and allow the retina to air dry for 5-10 s.
   NOTE: Failure to allow a brief air drying may randomly result in tissue-OCT separation during cryosectioning. Excessive air drying may lead to salt crystal-related tissue damage.
6. Apply OCT medium to cover the retina and use forceps to spread this layer evenly across the coverslip, preventing retinal flotation. Allow 30 s for OCT medium to penetrate the retina.
7. Flash-freeze the retina in OCT (Figure 5D) using either a metal plate indirectly cooled with liquid nitrogen vapor or by bringing it in direct contact with dry ice.
8. Store the retinas on dry ice if using immediately or in a -80 °C freezer in a plastic bag or container containing desiccants if storing it for the long term.

3. Cryosectioning

1. Set the cryostat chamber to -18 °C and stage to -12 °C. Place the retina in the chamber to equilibrate to temperature.
2. Create an OCT dome on the specimen chuck. If using the same model cryostat, ensure that this dome occupies the inner 2-3 annuli. Once frozen, trim the OCT dome to create a flat stage surface parallel to the blade on the chuck to retain laminar flatness (Figure 6A).
3. Create or note the orientation guide relating the chuck to the cryostat stage.
   NOTE: On the model cryostat we recommend, the orientation guide is a central line etched into the 12 o'clock position above the chuck holder, while the chuck itself has a triangular cut out on the outermost annuli. Together, we refer to these features as the orientation guide. If your cryostat does not have such features, we recommend using a permanent marker to make your own.
4. Prepare the retina for transfer using a razor blade by removing excess OCT surrounding the retina laterally and then, carefully separating the retina from the coverslip to make it mobile.
5. Remove the chuck from the stage and apply a small drop of liquid OCT to the prepared flat OCT stage surface.
6. Place the retina directly onto the liquid OCT. Using a prechilled (-18 °C) glass slide, press and hold the retina against the OCT stage for 10-30 s until frozen.
7. Apply additional OCT over the retina and stage to ensure complete bonding. Freeze on the cryostat's rapid freezing area. Incubate the retina/OCT stage in the cryostat for minimum 30 min to ensure complete bonding.
8. Begin sectioning:
   1. Carefully trim excess OCT until the retina becomes visible (Figure 6B). Use the razor blade to remove excess OCT from block edges.
      NOTE: Always retract the stage before resuming sectioning after any manual, radial trimming. For trimming slices en face, we recommend using the same cut thickness as will be used for desired sections; however, trimming thickness may be between 10 µm and 100 µm based on user preference.
   2. Continue trimming while inspecting sections for the presence of sclera. When the sclera is visible, adjust the stage angle as needed. Continue trimming until sections show equal sclera distribution, which ensures optimal flatness and maximum cloverleaf area in each section.
   3. Collect tissue sections (Figure 6C).
      NOTE: Optimal thickness will depend on the final assay being performed and must adhere to this downstream protocol. For example, if 10X Xenium is employed we recommend 20 µm-thick sections. Whereas for 10X Visium, 15 µm-thick sections are more appropriate.
   4. Use prechilled slides and manipulate the sections into position using two paintbrushes.
   5. Place a finger under the section until the OCT melts and tissue adheres (approximately 5 s).
   6. Return the slide to a cold surface for tissue re-freezing.
      NOTE: When collecting sections in small areas (e.g., for Xenium), multiple freeze-thaw cycles may be necessary. While this does not affect probe-based techniques, it may impact RNA quality for unbiased sequencing approaches.
9. Store the slides on dry ice for immediate use, at -20 °C for RNAscope the following day and at -80 °C for long-term storage.

4. Target detection and microscopy

NOTE: While the technique is compatible with other molecular profiling methods, we detail RNAscope as one validated example.

1. Sample preparation:
   1. Remove the slides from -80 °C storage and warm at -20 °C for 1 h.
      NOTE: Slides can be stored at -20 °C for up to 12 h prior to beginning the rest of this procedure but should not be left at -20 °C for too long to minimize RNA degradation.
2. Quickly heat the slides at 37 °C for 10 s.
3. Fix the tissue by immersing the slides in fresh 4% PFA for 30 min at room temperature.
4. Wash the slides thoroughly with 1x PBS.
5. Tissue dehydration through ethanol series: 50% ethanol for 5 min, 70% ethanol for 5 min, 100% ethanol for 5 min, fresh 100% ethanol for 5 min.
   NOTE: This is a critical quality control step for tissue mounting. If tissue detachment occurs during dehydration, it indicates OCT overlap from adjacent sections during mounting. Careful trimming of excess OCT during sectioning and mounting prevents this issue.
6. Barrier preparation:
   1. Air dry the slides on the benchtop for 5 min at room temperature.
   2. Using a hydrophobic barrier pen, draw a barrier around each tissue section. Allow the barrier to dry completely for 5 min at room temperature.
7. Molecular detection:
   1. Apply Protease IV treatment for tissue permeabilization.
   2. Proceed with the manufacturer's protocol for target probe hybridization (2 h at 40 °C), signal amplification steps (three sequential amplifiers), fluorophore detection (four sequential rounds for three-plex staining).
8. Imaging:
   1. Perform DAPI nuclear counterstaining prior to each cover glass mounting.
   2. Mount the coverglass using antifade mounting medium.
   3. Image using fluorescence microscopy with appropriate filter sets.
   4. For multiplex detection, capture the same field across all four rounds of detection for computational registration.
   5. Maintain consistent exposure settings between rounds for accurate signal comparison.
      NOTE: For complete details on RNAscope implementation, refer to manufacturer protocols (see the Table of Materials). For fully automated platforms like Xenium, follow all manufacturer protocol steps exactly as provided. For all protocols, anticipate that failure due to tissue detachment will most likely occur during an early dehydration or washing step, because of OCT overlap, as noted in this exemplar protocol.

Results

Successful implementation of this protocol produces intact en face retinal sections suitable for multiple molecular profiling approaches while maintaining spatial relationships across the retinal surface. We demonstrate the protocol's utility through a systematic validation approach, progressing from conventional immunohistochemistry through manual multiplex RNA detection to automated high-throughput molecular profiling.

The initial validation of our protocol employs two well-characte...

Discussion

The method presented here addresses a critical technical barrier in retinal spatial biology by enabling the production of thin en face sections suitable for modern molecular profiling technologies. The most critical steps in the protocol center around tissue handling during the initial dissection and the precise control of temperature and hydration during cryopreservation and sectioning. Successful implementation relies particularly on achieving proper tissue flattening before OCT embedding and maintaining consi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ames′ Medium	Sigma-Aldrich	A1420-10X1L	Specific brand not important
Beadsmith Thread Zap, Thread Burner	The Beadsmith Store	7.90524E+11	Alternative cauterizers may be used
Cryostat	Leica	cm1950	Important considerations is seperately controllable volume vs stage temperature.
Dry Ice	NA	NA	NA
Fine Science Tools Dumont #5 Forceps	Fine Science Tools 1129500	NC9889584	Specific brand not important
Fisher Healthcare Tissue-Plus O.C.T. Compound	Fisher Scientific	23-730-571	Specific brand not important
Fisherbrand High Precision Straight Slender Fine Point Tweezers/Forceps	Fisher Scientific	12-000-127	Specific brand not important
Fisherbrand Standard Dissecting Scissors	Fisher Scientific	08-951-20	Specific brand not important
Fisherbrand Superfrost Plus Microscope Slides	Fisher Scientific	22-037-246	Specific brand not important
Integra Miltex Noyes Iris Scissors	Integra Miltex 181510	12-460-278	Specific brand not important
Olympus IX81 spinning disk epifluorescence microscope			For Figure 7
RNAscope HiPlex	ACDBio
Rectangular Cover Glasses	Fisher Scientific	12-541-033	Specific brand not important
Sodium bicarbonate	Sigma-Aldrich	S6014-25G	Specific brand not important
Thermo Scientific Nunc Glass Bottom Dishes	Fisher Scientific	12-567-400	Specific brand not important
World Precision Instrument Precision Stereo Zoom Trinocular Microscope (IV) on Boom Stand	World Precision Instrument PZMTIVBS	50-253-8607	Specific brand not important
Zeiss Axio Imager.M2 epifluorescence microscope			Optical sectioning enabled (for Figure 8)