Preserving In Vivo Tissue Complexity in a Mice Olfactory Nervous System Model Using Whole-Mount Analysis for Studying Neural Regeneration

Summary

This protocol outlines a complete workflow for preserving the anatomical complexity of the mouse olfactory nervous system. Here we have detailed the complete workflow, including dissection, tissue preparation, and imaging. This protocol can be used to study the olfactory system under normal or abnormal conditions, including its natural repair conditions.

Abstract

In the era of three-dimensional (3D) imaging, standard immunohistochemistry has been recognized as highly limited in studies of pathology and complex human anatomy. By analyzing only narrow slices, larger structures and essential aspects of tissue architecture may lose critical contextual details, leading to incomplete or misleading interpretations. While two-dimensional (2D) sections have supported a basic understanding of histoarchitecture, they are insufficient for capturing the true three-dimensional reality of biological tissues.

Increasing numbers of studies across multiple anatomical regions now adopt techniques that isolate intact anatomical units - flat mount, whole-mount, or en face preparations - combined with high-resolution fluorescent microscopy. These approaches offer a more comprehensive visualization and interpretation of structural organization. This is particularly significant in the olfactory nervous system, where defining complex cellular mechanisms underlying its remarkable regenerative ability requires a full-thickness context. Thin sections risk missing key pathophysiological features that remain poorly defined.

Given these limitations and the knowledge gap surrounding the cellular interactions driving olfactory regeneration, developing a refined whole-mount technique capable of preserving full-thickness tissue complexity was essential. Entire olfactory mucosae were isolated as full-thickness samples, accompanied by an optimized antibody-penetration protocol. High-resolution imaging (EVIDENT SpinSR) and 3D reconstruction enabled accurate visualization of cellular interactions in vivo.

Introduction

The olfactory nervous system (ONS) represents a unique and well‑established model for studying mechanisms of neural regeneration, as it retains the unique capacity for neuronal turnover throughout life¹. However, the potential for clinical translation of this neuro-reparative ability remains limited due to significant gaps in our understanding of the behaviors of the cells in the olfactory system^2,3,4. The cellular responses and anatomical organization of ONS, including primary olfactory neurons, olfactory ensheathing cells (OECs), and perineural fibroblasts, have been described in several animal studies, relying on ultrathin (<2 µm) tissue sectioning combined with severe physical injury models applied to the rodent ONS^1,2. The animals in these studies were usually terminated at 10 days post-injury, providing a small window on just one specific moment of the extensive and more complex remodeling of such tissue during neural injury². These studies reported how, after axonal death, OECs and fibroblasts would persist as continuous channels following the injury.

While informative, due to the many limitations explained above, the utilized approach provided a low-resolution conceptual overview at a superficial level of the ONS, ultimately precluding a comprehensive assessment of the real cellular organization within the intact ONS³. As a result, critical spatial relationships may have been overlooked.

Preservation of three‑dimensional tissue structure is essential for accurate anatomical and cellular analysis and has been successfully applied across multiple organs and pathologies^{4,5,6,7,8,9,10,11,12}. Whole‑mount approaches have been used to map neural organization within the olfactory mucosa^13,14, however, visualization of the cellular interactions with a full-thickness ONS structure, including a preserved lamina propria, has not been done yet.

In this study, we establish a workflow using a mouse model with whole‑mount tissue processing, immunostaining, high‑resolution imaging, and three‑dimensional reconstruction, enabling detailed analysis of ONS architecture. This protocol was developed to fill the gap in existing published methods, where the protocol conserves and preserves a full-thickness olfactory mucosa, including the lamina propria, enabling the study of intercellular interactions. Notably, this refined tissue collection, and processing methodology can be applied and combined with various olfactory system models.

Protocol

All experiments containing animals and transgenically modified cells were conducted with the approval of the Griffith University Biosafety Committee (NLRD/003/2020_var3) and the Griffith University Animal Ethics Committee (MHIQ/04/23/AEC) in accordance with guidelines of the Australian Commonwealth Office of Gene Technology regulator (OGTR).

NOTE: A total of 15 adult laboratory mice (8–9 weeks) from both sexes were used in the study. Specifically, S100β-DsRed transgenic mice in which all OECs express the bright red protein DsRed under control of the S100β promoter. S100β is part of a wider calcium-binding protein family, and it is commonly expressed in glial cells in the central nervous system, such as OECs. This model was optimal for the purposes of these experiments allowed easily localization of the target cell populations using endogenous fluorescence signals. The S100β-DsRed transgenic mice were previously generated, and it has been shown that the use of these transgenic mice provides improved detection of OECs compared to immunohistochemistry for fixed tissue analyses^15,16,17,18.

1. Head collection and tissue fixation

1. Euthanize the animals with rising CO₂ inhalation at experimental timepoints.
2. Ensure death is confirmed by the absence of corneal and pedal withdrawal reflexes, followed by cervical dislocation, or in accordance with the approved SOP.
3. Remove the entire head and jaws with scissors.
4. Remove the skin from the skull and remove the cartilage at the tip of the nose to expose the nasal cavity and ensure efficient penetration of the fixative agent.
   NOTE: This step of cutting the most frontal part of the nose cartilage of the animal was observed to be crucial to allow the correct fixation of the tissue within the nasal cavity.
5. Quickly rinse the heads in PBS 1× to remove excess blood.
6. Fix the tissue by overnight (12–18 h) submersion in 4% paraformaldehyde, for at least 6 h (sufficient time to fix the tissue of interest) at 4 °C on a roller.
   CAUTION: Paraformaldehyde is a hazardous chemical that releases formaldehyde vapor. It is toxic by inhalation, skin contact, and ingestion; causes eye, skin, and respiratory irritation; and is a suspected carcinogen. Handle only in a fume hood while wearing appropriate PPE, including gloves, a laboratory coat, and eye protection. Safe disposal of hazardous chemicals and biological waste generated during the protocol must be done in accordance with institutional and local regulations.
   ​NOTE: During optimizations, we observed that 6 h was enough to fix the tissue of interest; however, more than 12 h was likely to cause over-fixation and degradation of the mucosal structures.
7. Remove heads from fixative agent and wash them for 3 h in PBS 1× at room temperature (20–25 °C) on a roller.
8. Store heads in fresh PBS at 4 °C and collect the whole-mount mucosal tissue within 48 h

2. Whole-mount tissue collection

1. Perform tissue isolation under a microscope.
2. Cut heads in half using a scalpel, following the mid-sagittal line.
3. Expose the interior of the nasal cavity (Figure 1). Identify the anatomical regions and collect the mucosal tissue lining the entire anatomical section (Figure 1B).
4. Using a bone cutter, gently break and remove the septal bone covering part of the mucosal cavity.
   NOTE: See Figure 2A; This step is crucial to expose the concave cavity underneath the bone where the axon bundles run through the dorsal wall (Figure 1B).
5. Once this area is exposed, use curved forceps to gently detach and scoop out the tissue from the concave dorsal wall. Lift the tissue from the underlying bone and gently peel the remaining olfactory mucosa to collect the intact whole-mount preparation.
   NOTE: Extreme caution was required at this stage to avoid tearing the tissue.
6. Collect the tissue of interest from both right and left nostrils: the mucosa covers the septum, the dorsal left and right walls, and the cribriform plate of the nasal cavity (Figure 1A, B). Collect tissue from both nostrils, including the septum, dorsal walls, and cribriform plate, to isolate the anatomical region containing the primary olfactory structures (Figure 2C).
7. Wash the collected tissues in PBS 1× and remove any unwanted tissues, including bone fragments. Store the tissues at 4 °C in PBS 1× for future immunohistochemistry analysis.
   NOTE: If the tissue will undergo long-term storage, it is advised to store it in a solution of PBS 1× and 0.001% (W/V) Sodium Azide, at 4 °C.

3. Optimized staining protocol

NOTE: Commonly utilized staining protocols are usually optimized for thin-medium thickness sections (e.g., up to 50 µm). Hence, the protocol needed to be adapted to the much thicker tissue.
​A crucial step was observed to involve keeping the tissue floating inside Eppendorf tubes during the entire staining process and placing the tubes on a roller while the reagents are penetrating. Therefore, the staining steps were not performed with the tissue on an imaging slide. The tissue was stained first while floating and subsequently mounted on imaging slides. These details ensured that the tissue would not fold, and the reagents and antibodies could penetrate the entire region.

1. Remove tissues from storage and place them into an Eppendorf tube, with blocking solution (solution of use will be staining specific).
   ​NOTE: Blocking solution: 3% BSA, 0.3% Triton X-100 in PBS. Another important adjustment was the time that the tissue was subjected to each reagent and antibody cocktail.
2. Keep tissue in blocking solution for 3 h at room temperature on roller.
   NOTE: It was observed that it was not necessary to increase the concentration of the selected antibodies and staining reagents (Table 1) if the time of exposure to them was increased compared to the standard thin section immunohistostaining approach. This minimized the usage of expensive antibody reagents as well. Incubations are done in a 1.5 mL Eppendorf tube with the lid-retainer cut off so it can roll on a roller. Reagent volume was 500 µL.
3. Incubate for 72 h at 4°Cwith primary antibodies on roller.
4. Wash tissue twice (1.5 h each wash) using PBS 1× at room temperature on roller.
5. Incubate tissue with secondary antibody (wrapped in aluminum foil) and nuclei staining for a minimum of 4 h at room temperature on a roller first and then move overnight (12–18 h) at 4 °C on a roller.
6. Wash tissue twice (1.5 h each time) using PBS 1× at room temperature on roller.

4. Slide preparation

1. Gently remove tissue from the Eppendorf tube
2. Under a microscope, place the tissue on a slide and ensure no folds in the tissue are present.
   NOTE: It is possible to identify under the microscope, without any fluorescence, the orientation of the tissue. The lamina propria side of the mucosal tissue should be placed on the slide with the epithelium facing the glass slide and the lamina propria facing the coverslip. The lamina propria can be identified by the presence of thick bundles of nerves, which are visible without staining. Having the orientation with the lamina propria facing the coverslip side, and the epithelium side lying on the slide glass, will facilitate the staining as well as imaging process.
3. Coverslip slides using fluorescent mounting medium and seal the slides.
4. Slides can be stored at 4 °C, after drying overnight (12–18 h) in the dark at room temperature

5. Image Acquisition and Post-Processing

1. Acquisition of low-magnification immunofluorescent images
   1. Perform automatic stitching during image acquisition using linear alpha-blending.
      ​NOTE: The number of z-stack frames should be adjusted according to the thickness of the specimen to ensure complete coverage of the tissue volume.
   2. Capture adjacent fields of view with 15% overlap. Acquire tiled images using a meander acquisition pattern.
   3. Adjust the total number of optical sections according to the thickness of the tissue.
   4. Set the image calibration to 215.594 nm/pixel in both the X and Y axes. Acquire images as z-stacks using a z-step size of 0.8 µm.
   5. Place the mounted whole-mount tissue on the confocal microscope stage. Acquire overview images using a 30x objective lens (NA 1.04).
2. Acquisition of high-magnification immunofluorescent images.
   1. Acquire detailed images using a 60× objective lens (NA 1.5). Set the image calibration to 107.923 nm/pixel in both the X and Y axes. Acquire z-stack images using a z-step size of 0.41 µm.
   2. Adjust the total number of optical sections according to tissue thickness. And capture images from a single field of view.
      ​NOTE: No image stitching is required for 60× acquisitions because each dataset consists of a single field of view.
3. Acquisition of histological reference images
   1. Image HE-stained mid-sagittal mouse head sections using a slide-scanning microscope under brightfield illumination.
   2. Acquire images using a 40× objective lens (NA 0.95) and a 0.63x camera magnification, resulting in an effective magnification of 25.2×.
   3. Confirm a pixel calibration of 136.905 nm/pixel in both the X and Y axes. And acquire the image as a single focal plane.
   4. Stitch tiled images automatically using a meander acquisition pattern and linear alpha-blending.
4. Acquisition of overview immunofluorescent reference images
   1. Acquire complementary overview immunofluorescent images using a 10x objective lens.
   2. Capture the specimen as a single z-slice by using 0% tile overlap during image acquisition.
   3. Set the image calibration to 647.249 nm/pixel in both the X and Y axes. And, generate an overview image encompassing the entire anatomical region of interest.
5. Image processing and generation of final figures.
   1. Export acquired image datasets to image analysis software. Generate maximum intensity projections (MIPs) from the z-stack datasets.
   2. Select regions of interest (ROIs) for subsequent analysis and figure preparation.
   3. Combine overview immunofluorescent images with corresponding HE images to generate anatomical reference figures. Then, verify alignment and anatomical correspondence between fluorescence and histological datasets before figure preparation.
      NOTE: All image stitching should be performed automatically during image acquisition using predetermined microscope settings and should not require additional post-processing. In our case, the 3D rendering was generated using the Imaris software: the surface rendering plugin, based on threshold intensity for each acquired channel, was used to produce the 3D reconstruction. Ultimately, final images were compiled on Adobe Illustrator.

Results

Our whole-mount approach presented here enabled the isolation and preservation of the olfactory mucosa in its entirety. In our case, our focus was on understanding cellular and anatomical changes on both macro (full anatomical view) and micro (cellular level) scales.

The whole-mount approach enables us to collect both epithelial and lamina propria compartments and preserve the spatial relationships between axon bundles, olfactory ensheathing cells (OECs), and fibroblasts – our cells of interest. This approach allowed the acquisition of large, continuous datasets that captured the full thickness and broad surface area of the mucosal tissue (Figure 3) and enabled us to capture details of the intricate cellular structure and interaction (Figure 3), and how these would change between intact (n = 3) (Figure 3A, B) and damaged conditions (n = 12) (Figure 3C, D).

By combining this optimized tissue handling and staining procedure with high‑resolution confocal microscopy, we were able to obtain images of wider mucosal regions with great and precise cellular‑level detail across the entire depth of the tissue. Z‑stack acquisition across multiple adjacent regions allowed reconstruction of extended anatomical domains without compromising resolution or signal integrity (Figure 4A–F). Due to the high-resolution cellular details, we were also able to use the acquired Z-stacks of the full tissue thickness (Figure 4G) to realize rendered images (Figure 4H) to further understand the tissue micro-architecture.

Reliable determination of tissue orientation prior to mounting and imaging was consistently achieved without the need for fluorescent labeling. The lamina propria side was readily identifiable under brightfield microscopy by the presence of thick nerve bundles, clearly distinguishable from the epithelial layer. Standardizing sample orientation by positioning the lamina propria adjacent to the coverslip and the epithelial surface against the glass slide improved imaging consistency across samples and facilitated downstream three‑dimensional reconstruction. Antibody penetration can be determined by analyzing the associated signal observed throughout the thickness of the tissue. It is usually evident by either an absence of signal or a signal that progressively decreases upon approaching deeper into the tissue.

Three‑dimensional rendering performed in Imaris enabled clear visualization of complex cellular networks and axonal organization within the intact mucosa. Surface‑based rendering, applied through channel‑specific intensity thresholding, generated accurate 3D representations of the tissue architecture while preserving fine structural details.

For staining and visualization, expected outcomes would include clear visualization of the target staining signal without any abnormal loss of the signal or deterioration of the signal throughout the thickness of the tissues. Inability to visualize a known stained target on microscopy should be considered an unsuccessful outcome.

Figure 1. Representation of the anatomical region with overlay of the mucosa whole-mount tissue. (A–C). Sagittal view of the inside of the left nostril (Hematoxylin and Eosin, HE staining) of the mouse (A) with overlay of mucosa whole-mount staining from the same region (for anatomical orientation) (B) displaying the orientation and position of axon bundles in green with cell nuclei in blue. This technique enabled the preservation of the entire anatomical structure: it is possible to clearly distinguish the primary olfactory nerve (ON) and accessory (vomeronasal nerves, VNN) axon bundles as well as follow their path from the septum to the olfactory bulb (OB), through the cribriform plate (B, C). Tissue displayed here was stained with Hoechst for nuclei (blue) and β-III-Tubulin for axon bundles (green). Representative images from n = 15 independent preparations. Please click here to view a larger version of this figure.

Figure 2. Macro and schematic views of the olfactory system. (A) Using a bone cutter, following the red dotted line, the septal bone is removed (yellow region). (B) Once the underlying concave nostril cavity is exposed (brown), the entire mucosal tissue expanding from this region to the respiratory and olfactory mucosa can be collected. (C) Graphical representation to illustrate the continuity of these structures extending from the olfactory epithelium through to the olfactory bulb. Please click here to view a larger version of this figure.

Figure 3. Extended view of the anatomical area obtained with a whole-mount collection and staining protocol. (A–D). Representative images from healthy and damaged mucosae are shown. From top to bottom, intact mucosa from healthy tissue (n = 3) and damaged mucosa (n = 12). In the healthy tissue (A), OECs with their elongated bipolar morphologies (A, left) wrap the axon bundles (A, middle), and fibroblasts form the outer layer of these tunnel-like structures (A, right, white arrow). The three selected ROIs reflect these details (B). In the damaged mucosa, OECs display both bipolar and round morphologies (C, D, left, white arrows). Axon bundles appear less uniform and solid (C, D, middle, white arrows). Fibroblasts appear widespread throughout the entire region (C, D, right, white arrows). Scale bar = 100 µm. Please click here to view a larger version of this figure.

Figure 4. Three-Dimensional fluorescent imaging of mucosal whole mounts. (A–H). Low-power images of the cells (A–F) and a high-power image (G) were obtained from image stacks of multiple consecutive regions using a SpinSR microscope. Olfactory ensheathing cells (OECs) are shown in red, fibroblasts in magenta, axons in green, and nuclei in blue. Arrows indicate different OEC morphologies (A) and an outer layer of fibroblasts around axon bundles (B and C). D–F have arrows highlighting similarities in the fibroblasts’ healthy structures. These images highlighted the spatial organization of the cellular components, which were then rendered using Imaris software (H). Scale bar = 50 µm. Please click here to view a larger version of this figure.

Reagent	Target	Dilution	Incubation conditions	Rationale
Primary antibody
β-III-Tubulin (Rabbit)	Axons	1:300	For 72 h at 4 °C, on roller	to allow full thickness penetration
PDGFR-β (Rat)	Fibroblasts	1:100
Secondary antibody
Alexa 488	Rabbit primary antibodies	1:250	Wrapped in aluminum foil, for a minimum of 4 h at room temperature on a roller, then overnight at 4 °C on a roller.	Aluminium foil prevents photobleaching, The duration, temperature and roller-induced agitation enables good penetration
Alexa 647	Rat primary antibodies	1:500
Nuclei
Hoechst	Cell nuclei	1:2500

Table 1: List of antibodies with dilutions optimized for this protocol. This table lists the staining reagents with their targets, dilutions, and rationale for the incubation conditions.

Discussion

The last few decades have slowly witnessed the start of the three-dimensional era, with many innovative approaches developing in the scientific field. For instance, three-dimensional reconstruction approaches, made possible by the development of complex whole‑mount imaging methods paired with high‑resolution fluorescent microscopes, have facilitated a better understanding of several full‑thickness tissue histoarchitectures. These three‑dimensional rendering approaches have been used to define several human body structures, such as vessels, glands, and common tumors^4,5,6,7,8,9. Moreover, information on the differences between pathological and normal structures has been collected, for example, in full mouse brains^10,11,12. This approach has been identified as a crucial step towards a better understanding of the various biological systems, facilitating a deeper comprehension of cellular circuits, their structural and functional spatial organization, and their potential contribution to human pathologies.

A downside of the different combinations of three‑dimensional tissue reconstruction and imaging is that most approaches involve tissue-clearing steps. Although this method is useful for many tissues, it is not appropriate for all tissue types^8,9,19. In fact, clearing procedures involve several washing steps, which could potentially damage targeted epitopes and influence endogenous signals²⁰.

The above‑described approaches have been used to define many important cellular structures within different systems. However, the olfactory system and its structures of interest have a minimal presence of bony components from which they could be easily isolated; therefore, whole‑mount approaches that do not include clearing steps have been trialed in recent years^13,14.

These studies recognized the importance of better defining the organization of neuronal structures in the olfactory system. Accordingly, their main focus was to provide a clear topographic map of the olfactory sensory neurons’ distribution based on the expression of different markers¹⁴ as well as to reveal novel insights into their peculiar cilia¹³.

The whole‑mount technique developed and presented here enables the collection and preservation of the mucosal tissue in its entirety, opening the possibility of gaining crucial insights into the anatomical and physiological structure of this environment, with a specific focus, in our case, on the relationship between OECs, fibroblasts, and axon bundles. Pairing this refined isolation and staining technique with high‑resolution confocal imaging enabled the collection of large images that reflect the structure of the entire target region and its resident cells. Specifically, our refined technique offers two distinctive advantages where the entire mucosa is harvested and processed, including the lamina propria architecture¹³, and the gentler tissue preparation method helps preserve the macro-scale anatomy as well as inter-cellular interactions (such as olfactory ensheathing cells, axons, and fibroblasts) compared to the other flat-mount protocols¹⁴. This is also why this protocol does not include validation against thin section staining.

As mentioned above, whole‑mount techniques usually involve long tissue‑processing periods and the use of specific, expensive reagents. This method was developed with a focus on minimal tissue processing to minimize both the timeframe and the risk of damaging important cellular structures. The optimized method, therefore, represents a quick process that can be performed using reagents commonly employed for immunohistochemistry on thin sections, providing a valid alternative to approaches that require longer processing times and are not appropriate for boneless structures such as mucosal tissue.

Image acquisition and post‑processing can be performed using any confocal microscopy platform capable of acquiring high‑resolution z‑stacks from thick tissue samples, with sufficient laser penetration to image through the mucosae of interest (refer to the methods section for more details on our acquisition and image processing settings).

With appropriate modifications, we believe that this protocol can be translated and applied to other common rodent models. With some tissue‑specific modifications, this method can be applied to several different contexts, including the study of mucosal and submucosal tissues from the respiratory, gastrointestinal tract (GIT); the investigation of tissue organization and multi‑system interactions within the nervous system; the analysis of inflammatory processes and mucosa‑associated lymphoid tissues (MALTs); as well as the exploration of different multisystem axes, such as the gut–brain axis.

The main strength of this protocol lies in the methodological refinements that allow for high‑quality three‑dimensional imaging while preserving tissue integrity but also ensuring a relatively fast approach compared to other whole-mount methodologies previously trialed in the field.

The successful implementation of this approach was strongly dependent on a deep anatomical understanding of the tissue of interest. In fact, in our case, knowing the expected localization of the axon bundles was essential to minimize the invasiveness of the collection procedure once the inside of the skull was exposed. This anatomical knowledge was crucial in being able to quickly localize the region and tissue of interest and preserving the overall structure of the mucosa, highlighting how successful tissue processing is intrinsically linked to an informed collection strategy rather than post‑processing steps alone.

As mentioned in the methods section in several notes, multiple technical adjustments were critical to the development and optimization of this method. Maintaining the tissue floating freely within Eppendorf tubes throughout the entire staining process, combined with continuous rotation during reagent and antibody incubation, represented a key step from more static whole‑mount and thin-section protocols. This refinement prevented tissue folding while also ensuring homogeneous reagent penetration across the entire sample, thus improving staining consistency without the need for unnecessary clearing steps. In addition, careful optimization of the incubation duration for each reagent and antibody cocktail proved essential. This last step might need to be further adjusted based on the type of antibody chosen.

Together, these specific factors resulted in enhanced reproducibility while reducing overall processing time and harsher sample handling.

Another important methodological step was the ability to reliably determine tissue orientation prior to mounting and imaging without the need for fluorescent labeling. The lamina propria side of the mucosal tissue could be readily identified under brightfield microscopy by the presence of thick nerve bundles, clearly distinguishable by the surrounding epithelial tissue. Standardizing sample orientation by positioning the lamina propria facing the coverslip side and the epithelial layer resting on the glass slide was found to facilitate image acquisition quality. This simple but effective refinement improved imaging consistency across samples and contributed to the quality of the three‑dimensional reconstructions obtained.

There are some limitations to consider, however. This protocol has not been tested on many common mouse strains or other rodent species. Similarly, the optimizations were conducted under the specific experimental conditions as described above and with the listed antibody panels alone. While the protocol should work with most mouse and rat strains, several different antibodies, and any experimental models that allow for mucosae to be harvested in full thickness, further careful optimization may be required. Another notable limitation is that a single whole-mount section can only be stained with one antibody panel, unlike thin-sectioned tissues, where each section can be stained with individual antibody panel and therefore, multiple different panels of antibodies can be tested on thin-sectioned tissues.

Overall, these methodological innovations demonstrate how relatively straightforward adjustments to tissue handling, staining conditions, and sample orientation can substantially improve whole‑mount imaging outcomes. By avoiding extensive tissue processing and clearing steps, this approach offers a practical and adaptable alternative for the study of delicate, boneless tissues, while still enabling the acquisition of detailed, large‑scale three‑dimensional data.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Bovine Serum Albumin (BSA)	Sigma-Aldrich	A3294-50G	Immunohistochemistry
Adobe Illustrator		version 28.5
Anti-beta III Tubulin antibody - Neuronal Marker	Abcam	ab18207
Anti-PDGFR beta antibody [APB5]	Abcam	ab91066
bisBenzimide H 33342 trihydrochloride	Sigma Aldrich	14533
COVERSLIP 24X50MM NO. 1.5 100PCS	Bio-strategy	EPBRCS245015GP	Immunohistochemistry
Donkey anti-Rabbit IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluo 488	Thermo Fisher	A-21206
Dumont #15A Bone Cutter - Epoxy Coated	Daniels Health Laboratory Products P/L	11215-02	Tissue dissection
Dumont #7 Forceps - Biologie/Dumostar	Daniels Health Laboratory Products P/L	11297-10	Tissue dissection
Goat anti-Rat IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor™ 647	Thermo Fisher	A-21247
Imaris software	Imaris	version 9.5.0	Imaging
Mayo dissecting scissors	ProSciTech	420SS	Tissue dissection
Paraformaldehyde	Sigma-Aldrich	P6148-500G	Tissue Processing
Phosphate buffered saline, sterile PBS Tablets (PBS)	Thermo Fisher Scientific	18912014	Tissue Processing
ProLong Glass Antifade Mountant	Thermo Fisher Scientific	P36980	Immunohistochemistry
Scalpel blade #23	Scalpel Blades Swann Morton	COS240B	Tissue dissection
Scalpel handle #4	ProSciTech	T134	Tissue dissection
Sodium Azide	Sigma-Aldrich	S2002-25G	Tissue Processing
Sucrose	Chem-Supply	57-50-1	Tissue Processing
TOMO Adhesion microscope slides	ProSciTech	GEMS63705-09	Immunohistochemistry
Triton X-100	Sigma-Aldrich	X100-100mL	Immunohistochemistry