Multiplex Fluorescent mRNA In Situ Hybridization of Developing Craniofacial Tissues in FFPE Mouse Samples

Summary

Conventional in situ hybridization struggles to visualize mRNA in craniofacial tissues due to complex architecture, high background, limited probe specificity, and sensitivity. This protocol describes a multiplex fluorescent RNA in situ hybridization method for sensitive, specific detection of multiple mRNA targets in FFPE mouse embryonic craniofacial tissues, overcoming previous limitations.

Abstract

Craniofacial development depends on the coordinated activity of diverse, specialized cell populations that interact within intricate tissue architectures to form and maintain normal tissue structure and function. Understanding how these cells communicate with each other and with their local microenvironment is essential for defining the molecular mechanisms that regulate their formation and homeostasis. Disruption of these processes can lead to congenital craniofacial conditions, including tooth agenesis, cleft palate, and cleft lip, either alone or as part of syndromic disorders. While quantitative techniques like RT-PCR offer high sensitivity and dynamic range, they require RNA extraction from homogenized tissue, resulting in a loss of spatial and cellular context, an especially critical limitation for heterogeneous craniofacial tissues, where the cellular origin of transcripts cannot be resolved. Multiplex fluorescent RNA in situ hybridization overcomes this limitation by enabling sensitive and specific detection of multiple mRNA transcripts while maintaining tissue architecture. This advanced approach allows precise localization of gene expression at the cellular level, providing valuable insight into the spatial regulation of developmental processes and disease mechanisms in craniofacial biology.

Introduction

Real-time polymerase chain reaction (RT-PCR) is the gold standard for gene expression analysis due to its high sensitivity and broad dynamic range; however, it requires RNA extracted from homogenized tissues, resulting in loss of spatial and cellular context¹. This limitation is particularly problematic for heterogeneous tissues such as craniofacial tissues during embryonic development, where the cellular source of detected transcripts cannot be resolved. RNA in situ hybridization (ISH), a technique pioneered in 1968 by Joseph Gall and Mary Lou Pardue, overcomes this limitation by enabling the localization of mRNA expression within intact tissue architecture². Nevertheless, conventional RNA ISH is technically demanding and highly dependent on optimal tissue fixation and sample preparation to preserve RNA integrity³. These challenges are exacerbated in highly calcified tissues, which require decalcification procedures that often involve harsh chemicals or prolonged processing, leading to RNA degradation. Consequently, RNA ISH in craniofacial tissues, which comprise complex mixtures of cell types, necessitates careful optimization and technical expertise, is time-intensive, and carries a substantial risk of suboptimal staining. The challenge of visualizing mRNA expression is even more pronounced in dental tissues, which possess a highly complex structure and composition. Teeth are composed of mineralized components such as enamel, dentin, cementum, and surrounding bone, alongside softer tissues including the pulp and periodontal ligament. These tissues are populated by a diverse array of cell types that play crucial roles in tooth development⁴. These features require careful standardization to ensure reliable signal detection. As a result, the described protocol is particularly well suited for investigating gene expression during developmental and postnatal stages of tooth formation.

Multiple fluorescence in situ hybridization (FISH)-based techniques exist for spatial RNA detection, including probe-based amplified RNA FISH^5,6,7,8,9, hybridization chain reaction FISH⁶, single-molecule RNA FISH¹⁰, signal amplification by exchange reaction FISH⁵, and highly multiplexed FISH platforms like MERFISH¹¹ and seqFISH^12,13. Among these, multiplex fluorescent RNA ISH^7,8,9 is widely used due to its high sensitivity in detecting a single RNA-molecule resulting from the double Z-probe design, in which two adjacent probes must hybridize to the target sequence to initiate signal amplification. This dual recognition design greatly improves specificity and reduces off-target signal. The method supports multiplex detection of 2–4 targets (expandable up to 12 or more targets using similar techniques), preserves tissue morphology, and is compatible with formalin-fixed paraffin-embedded (FFPE)^7,8,9, fresh-frozen, and selected calcified tissues, making it suitable for complex samples such as those encountered in craniofacial and dental development. The method is standardized, reproducible, supported by well-established probe libraries, and widely validated, ensuring accessible and reliable results across various laboratories.

Multiplex fluorescent RNA ISH employs multiple probes targeting distinct regions of the same transcript combined with enhanced signal amplification, enabling highly sensitive and specific detection even for low-abundance or partially degraded mRNAs^14,15. While multiplex fluorescent RNA ISH has been successfully applied to decalcified bone samples¹⁶, and its use in complex craniofacial tissues has previously been reported^7,8,9; however, a detailed step-by-step protocol describing its application in this context has not yet been reported.

Craniofacial structures comprise intricate tissue architectures with diverse cell types, including osteoblasts, osteoclasts, osteocytes, ameloblasts, odontoblasts, fibroblasts, stem cells, immune cells, vasculature, and sensory neurons, whose spatially coordinated interactions are essential for tissue formation, function, regeneration, and immunological protection⁴. Studying localized gene expressions in these tissues is therefore critical, and multiplex fluorescent RNA ISH represents a leading approach for such analyses.

In this study, we described how multiplex fluorescent RNA ISH based on RNAscope probes and technology could be applied to study complex craniofacial tissues (Figure 1). Furthermore, we established an optimized FFPE sectioning workflow that preserves RNA integrity and enables high-quality, spatially resolved gene expression analysis.

Protocol

All animal procedures were approved by the National Institutes of Health, National Institute of Child Health and Human Development Animal Care and Use Committee (IACUC), under Animal Study Protocol #24-031.

1. Timed mating and embryo tissue collection

1. Set up timed matings.
   1. Pair a healthy, fertile male and female Mus musculus in the late afternoon. The following morning, check for the presence of a vaginal plug. Designate the day a plug is observed as embryonic day (E) 0.5.
2. Euthanize the pregnant female.
   1. At the desired developmental stage, euthanize the pregnant female mouse via CO₂ inhalation followed by cervical dislocation, in accordance with institutional animal care guidelines.
   2. Position the mouse on its back on a sterile absorbent surface. Disinfect the abdominal area with 70% ethanol along the intended incision site.
3. Expose and isolate the uterine horns.
   1. Make a midline abdominal incision with sterile surgical scissors, then open the peritoneal wall with microsurgical scissors to expose the uterine horns.
   2. Use blunt forceps to gently exteriorize the uterine chain. Release the uterine tissue along the myometrium and cut the oviduct and the median uterine horn attachment to isolate the embryos.
4. Transfer whole embryos.
   1. Immediately transfer the uterine chain containing embryos to a 10 cm Petri dish containing ice-cold, sterile 1× PBS (pH 7.4).
5. Dissect individual embryos.
   1. Carefully separate each embryo from the surrounding uterine, amniotic, and chorionic membranes.
   2. Transfer the cleaned embryos to a fresh 10 cm dish containing ice-cold, sterile PBS.
6. Dissect mouse embryonic heads.
   1. Using a sterile surgical blade, dissect the heads and rinse the dissected heads once in ice-cold PBS before further processing.
      NOTE: Collect and dissect the neonatal mice at the desired developmental stage in accordance with the animal protocol.

2. Fixation and processing of tissue

1. Fix the tissue.
   1. Immediately immerse the dissected tissue in 10% neutral buffered formalin and fix overnight at room temperature on a shaker.
      ​NOTE: Proper preservation of RNA is crucial for the success of in situ hybridization analyses. Numerous studies have shown that the standard approach—fixing tissue samples in 10% neutral buffered formalin for durations ranging from 6 to 48 hours, depending on the specific size and age of the tissue—produces reliable results for RNA preservation^16,17,18,19. For earlier embryonic stages (E12.5 and below), the penetration of fixative is rapid, and the fixation time should be reduced accordingly to prevent over-fixation. From embryonic day 17.5 (E17.5) onward, as well as in postnatal samples, penetration of the fixative can be less efficient due to the presence of skin on the head, which acts as a physical barrier. Hence, the skin should be removed from the whole head to allow efficient penetration of the fixation solution. Decalcification can be done in 0.5 M EDTA (pH 8.0) containing 0.5% methanol-free formaldehyde at 4 °C⁷. The duration of decalcification can be monitored by a quick microCT scan and may be adjusted depending on the degree of mineralization and the specific tissue type; for E18.5 embryos, 24–48 h is usually sufficient.
2. Rinse and process the tissue.
   1. Remove the fixative and rinse the samples in PBS. Transfer the tissue to 70% ethanol and store at 4 °C until further processing (1–4 weeks)²⁰. Perform tissue dehydration, clearing, and paraffin infiltration using an automated tissue processor according to the manufacturer’s instructions.
      ​NOTE: The typical tissue processing protocol is the following: 70% ethanol for 30 min, followed by 95% ethanol for 30 min (2 times), and 100% ethanol for 30 min (3 times) for dehydration; clearing in xylene for 1 h (3 times), and subsequently infiltration with paraffin wax at 60 °C for 1 h (3 times) before embedding in paraffin tissue blocks.
3. Embed the tissue.
   1. Embed the heads in paraffin and orient the samples in either the coronal or sagittal plane, depending on the experimental requirements.
   2. Transfer the embedded blocks to a paraffin block cold plate to allow the wax to harden for approximately 30 min, with timing adjusted as needed based on block size.
4. Store the FFPE blocks.
   1. Allow the paraffin to solidify completely. Take blocks out of molds, put them in labeled airtight containers or zip-lock bags, and store at 4 °C until ready to section.
      NOTE: FFPE blocks with well-preserved RNA can be stored for several years prior to sectioning without compromising tissue integrity.

3. Processing of FFPE tissue for multiplex fluorescent RNA ISH

1. Prepare an RNase-free workspace.
   1. Disinfect all work surfaces and instruments with 70% ethanol. Treat surfaces, tools, and gloves with an RNase-decontaminating solution to minimize RNA degradation.
   2. Set up the microtome and install a new blade. Set the clearance angle to 10°. Treat tweezers and brushes with RNase-decontaminating solution prior to use.
      ​NOTE: Always use a new blade for sectioning to obtain precise tissue slices. Blade angle significantly affects section quality, tissue integrity, thickness uniformity, and preservation of morphological features.
2. Prepare the water bath.
   1. Fill a flotation water bath with ultrapure laboratory-grade water.
   2. Adjust the temperature between 40–43 °C as needed, depending on embryonic stage and tissue characteristics.
3. Trim and mount the FFPE block.
   1. Trim excess paraffin from around the embedded tissue. Cool the FFPE block on ice and mount it onto the microtome cold stage.
4. Trim to the region of interest.
   1. Align the paraffin block face with the microtome blade and trim at 12 µm steps until the region of interest (e.g., the adjacent to the molar or incisor tooth) appears in coronal sections.
   2. Reposition the block and blade as needed for symmetrical, properly oriented whole-head coronal or sagittal sections.
5. Section the tissue.
   1. Once the region of interest is reached, cut 5–7 µm sections. Float the sections on the water bath to allow gentle relaxation and flattening before transferring them to slides.
6. Mount and dry the sections.
   1. Mount the sections onto positively charged glass slides. Examine the slides under a light microscope to confirm proper orientation and tissue integrity.
   2. Dry the slides on a slide warmer at 40 °C overnight prior to performing the multiplex fluorescent RNA ISH assay.

4. Day 1: Deparaffinization and rehydration

1. Bake the slides.
   1. Bake the slides with FFPE sections for 1 h at 60 °C.
      ​NOTE: If the slide is not used immediately, store it not baked at room temperature (RT) with desiccants for 1–4 weeks. Extended storage may result in RNA degradation.
2. Equilibrate the slides.
   1. Allow the FFPE tissue slide to equilibrate at room temperature (RT) for 10 min.
3. Deparaffinize and rehydrate the sections.
   1. Immerse the slides in xylene or a xylene substitute with gentle agitation for 5–10 min at RT. Repeat this step once to ensure complete removal of paraffin.
      NOTE: Work with xylene should be performed in a chemical fume hood. If a low-toxicity xylene substitute is used, it can be used outside of the chemical fume hood.
   2. Transfer the slides to 100% ethanol and incubate for 2 min at RT. Repeat once to remove residual xylene. Rehydrate the sections in 70% ethanol for 2 min.
   3. Rinse the sections 3 times with PBS. Then add a drop of PBS sufficient to fully cover the sections and immediately proceed to the next step.

5. Multiplex fluorescent RNA ISH on FFPE tissue sections

NOTE: This study followed the manufacturer’s instructions for performing the multiplex fluorescent RNA ISH on FFPE tissue sections using RNAscope technology²¹.

1. Day 1: Pretreatment and probe hybridization
   1. Draw a hydrophobic barrier.
      1. Using a barrier pen, draw a hydrophobic barrier around each tissue section. Allow the slides to air-dry for approximately 10 min at room temperature (RT).
      2. Ensure that tissue sections remain covered with PBS at all times to prevent drying.
         ​NOTE: Use a lint-free wipe (lens wipe) to dry the slide around the tissue while keeping the tissue hydrated. Ensure that the barrier pen ink does not come into contact with the tissue section.
   2. Perform pretreatment.
      1. Apply the custom pretreatment reagent to completely cover each section. Incubate the slides for 30 min at 40 °C in a humidified chamber placed inside a hybridization oven.
         NOTE: For all reagents supplied in dropper bottles, add sufficient drops to fully cover the section within the barrier. Ensure all reagents are brought to RT before use.
   3. Rinse the sections.
      1. Rinse the sections several times with PBS, then incubate in fresh PBS for 2 min at RT.
         NOTE: Use a 5 mL transfer pipette to wash each slide individually with PBS. During the 2-min wash, ensure the sections remain fully covered with PBS.
   4. Quench endogenous peroxidase activity.
      1. Apply 3% hydrogen peroxide to each section and incubate at RT for 10 min to block endogenous peroxidase activity.
   5. Repeat the wash.
      1. Wash the slides in PBS for 2 min at RT. Repeat twice.
   6. Prepare the probes.
      1. Pre-warm the probes at 40 °C for 10 min. Allow the probes to cool to RT prior to use.
   7. Prepare the probe mixture.
      1. The C1 probe is supplied as a 1× working solution; use this directly.
      2. In contrast, C2, C3, and C4 probes are provided as 50× concentrates; dilute these probes with the C1 probe solution. If the C1 probe is not used, dilute C2, C3, and/or C4 probes with the probe diluent supplied by the manufacturer.
   8. Hybridize the probes.
      1. Apply 30–40 µL of probe mixture to each tissue section, ensuring complete coverage. Incubate the slides for 2 h at 40 °C in a humidified chamber in the hybridization oven.
         NOTE: When learning the RNAscope technique, it is recommended to include both negative and positive control probes during test runs. This approach helps to validate the assay and confirm the quality of the procedure. A potential limitation to be aware of is that some manufacturer-provided positive control probes may not be expressed in the tissue of interest. To address this, cell-type-specific probes can be used. For example, Col1a1 is a reliable positive control for connective tissues. Staining for Col1a1 (or another gene with known high expression) can be performed as a first step to assess RNA preservation and tissue morphology in the samples. Once proficiency with the technique is achieved, negative controls are typically carried out by processing slides without probes but treating them with all other solutions. This method provides critical information regarding the background signal in all detection channels. The standard Multiplex V2 kit enables simultaneous staining and imaging of up to four different probes. When the gene expression levels are established through prior transcriptomics studies, it is advisable to assign genes with low expression to the Cy5 and Cy7 channels. Genes that are highly expressed should be placed in the GFP and RFP channels for optimal detection. In cases where none of the probes target genes with high expression, the GFP channel should remain unused. This allows the GFP channel to serve as a reference for background subtraction from the RFP channel, improving the accuracy of signal detection for low-expressing targets.
   9. Rinse with wash buffer.
      NOTE: Commercially available 20× Wash Buffer was diluted to 1× with RNase-free water prior to use.
      1. Wash the slides for 2 min at RT in the 1× Wash Buffer provided with the kit. Repeat twice.
   10. Incubate in 5x saline-sodium citrate (SSC) buffer.
       ​NOTE: Commercially available 20× SSC Buffer  (3 M NaCl, 0.3 M sodium citrate, pH 7.0) was diluted to 5× with RNase-free water prior to use.
       1. After washing, incubate the sections overnight in 5× SSC buffer at RT.
2. Day 2: Signal amplification, fluorescent labeling, and mounting
   1. Wash the slides.
      1. Wash the slides in 1× Wash Buffer for 2 min at room temperature (RT). Repeat twice.
   2. Perform the amplification steps.
      1. Apply Amplification 1 reagent to each section and incubate for 30 min at 40 °C in a humidified chamber.
      2. Rinse twice in 1× Wash Buffer for 2 min at RT.
      3. Apply Amplification 2 reagent and incubate for 30 min at 40 °C.
      4. Rinse twice in 1× Wash Buffer for 2 min at RT.
      5. Apply Amplification 3 reagent and incubate for 15 min at 40 °C.
         NOTE: All three amplification steps are required regardless of which channels (probes) are being developed.
      6. Rinse twice in 1× Wash Buffer for 2 min at RT.
   3. Develop horse radish peroxidase (HRP) and fluorophore.
      1. Calculate the volume of fluorophore needed (generally 100–200 µL per slide) and dilute the Opal fluorophore stocks.
         ​NOTE: The recommended dilution range for fluorophore is 1:300–1:1500. In these experiments, Opal Polaris 780 (Fluorophore reagent) was diluted 1:500 in antibody diluent, and all other dyes were diluted 1:1000 in TSA buffer. Minimize light exposure to the slides whenever possible.
      2. Apply HRP-C1 reagent and incubate for 15 min at 40 °C.
      3. Rinse twice in 1× Wash Buffer for 2 min at RT.
      4. Apply 30–40 µL of Opal dye (e.g., Opal 570 or Opal 690) to each section. Incubate for 30 min at 40 °C.
      5. Rinse twice in 1× Wash Buffer for 2 min at RT.
      6. Apply HRP Blocker and incubate for 15 min at 40 °C.
      7. Rinse twice in 1× Wash Buffer for 2 min at RT.
      8. Apply HRP-C2 or HRP-C3 reagent and incubate at 40 °C for 15 min.
      9. Rinse twice in 1× Wash Buffer for 2 min at RT.
      10. Apply 30–40 µL of the second Opal dye and incubate for 30 min at 40 °C.
      11. Rinse twice in 1× Wash Buffer for 2 min at RT.
      12. Apply HRP Blocker and incubate for 15 min at 40 °C.
      13. Rinse twice in 1× Wash Buffer for 2 min at RT.
      14. Apply HRP-C3 or HRP-C4 reagent and incubate for 15 min at 40 °C.
      15. Rinse twice in 1× Wash Buffer for 2 min at RT.
   4. Perform tyramide signal amplification-digoxigenin (TSA-DIG) and fluorophore labeling (If applicable).
      1. Apply 30–40 µL of diluted TSA-DIG reagent per section and incubate for 30 min at RT.
      2. Rinse twice in 1× Wash Buffer for 2 min at RT.
      3. Apply blocking reagent and incubate for 15 min at 40 °C.
      4. Rinse twice in 1× Wash Buffer for 2 min at RT.
      5. Apply 30–40 μL of diluted fluorophore reagent and incubate for 30 min at 40 °C.
      6. Rinse twice in 1× Wash Buffer for 2 min at RT.
   5. Perform counterstaining and mounting.
      1. Apply 4′,6-diamidino-2-phenylindole (DAPI) and incubate for 5 min at RT in the dark.
      2. Rinse DAPI and promptly mount the sections using anti-fade mountant, apply the coverslip gently, and avoid trapping air bubbles. Select a coverslip size that appropriately matches the tissue coverage.
      3. Dry the slides for 2 h at RT in the dark, then keep the slides at 4 °C for long-term storage.
         ​NOTE: Fluorescence can be preserved for up to a year if stored at 4 °C in the darkness with proper anti-fade mounting.
   6. Perform imaging.
      1. Perform imaging using a multichannel fluorescence microscope with LED illumination and filters for spectral separation (Figure 2A). Use the excitation and emission wavelengths as follows: DAPI (353/465 nm), FITC (495/519 nm), Cy3 (548/561 nm), Cy5 (650/673 nm), and Cy7 (747/773 nm). Specific filters minimize spectral overlaps and distinguish blue, green, orange, red, and near-infrared signals.
      2. Acquire all images using identical exposure and illumination settings across samples, ensuring consistency and enabling reliable comparative analysis.

Results

Using the described protocol (Figure 1), we demonstrate multiplex fluorescent mRNA ISH transcript detection in a mouse whole head imaged on a Zeiss microscope platform (Figure 2B). Specific image views can be selected using the side tab (yellow box), while multiple channels can be visualized simultaneously using the channel tab (red box). Brightness, gamma, and contrast of the live image can be directly adjusted with the controls under the display tab (green box...

Discussion

In situ hybridization has been a foundational tool for studying gene expression for more than four decades and has undergone continuous refinement. However, conventional ISH approaches are often limited by low sensitivity, high background signal, and lengthy, labor-intensive procedures. These constraints have restricted their broader application, particularly in complex craniofacial tissues where precise spatial resolution and detection of low-abundance transcripts are essential. Multiplex fluorescent RNA ISH ov...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1×PBS	Thermo Fischer	10010023	Use  to perform washes during the workflow
20×SSC	Thermo Fischer	J60839.K2	Use to make the 1´ Wash buffer
50 mL conical tubes (Ambion) RNAse free	Thermo Fischer	AM12502	Use to store samples in different solutions
Advanced orbital shaker	VWR	6683-470	Use to shake tissues in fixation solution during incubation
Alcohol, 70%, Fisherbrand, HistoPrep	Fisher Scientific	HC-1000-1GL	Use to clean and disinfect all the work space
Antibody Diluent	Akoya Biosciences	ARD1001EA	Diluent for TSA-DIG
Automated vacuum tissue processor	Leica Biosystems	ASP300S	Use to clear, dehydration, rehydration and wax infiltration of samples
Cover Glass Thickness 1.5, 25 mm x 25 mm	Corning	2850-25	Use for mounting of slide in Visium HD workflow
Custom Pretreatment Reagent	ACD, Bio-Techne	300040	To optimize tissue permeabilization and target accessibility
EDTA	Millipore Sigma	03690	Use to decalcify mineralized tissues
Ethanol 100%	AeroBase Group	6505001050000	Use to remove xyline and rehydrate samples during processing
HistoCore Water Bath	Leica Biosystems	HIS2326	Use to float the sections at 40-43 °C to remove wrinkles from FFPE sections
HybEZ Humidifying paper	ACD, Bio-Techne	310015	Use inside the HybEZ oven to maintain humidity and prevent tissue sections from drying during hybridization
HybEZ Oven, Tray, and Rack	ACD, Bio-Techne	310011, 310012, 310014	Use to provide controlled temperature, humidity, and proper slide positioning during hybridization and incubation steps
ImmEdge Hydrophobic Barrier Pen	Vector Laboratory	H-4000	Use to draw hydrophobic boundaries on slides to confine reagents to the tissue section during staining procedures.
Low-Profile Disposable Blades DB80LX	Leica Biosystems	14035843496	Use to section FFPE blocks
Neutral Buffered Formalin 10%	Azer Scientific	NBF-4-G	Use to fix the tissues
OPAL 520 REAGENT PACK	Akoya Biosciences, Inc.	FP1487001KT	To amplify and detect target RNA signals with fluorescent labeling.
OPAL 570 REAGENT PACK	Akoya Biosciences, Inc.	FP1488001KT	To amplify and detect target RNA signals with fluorescent labeling.
OPAL 620 REAGENT PACK	Akoya Biosciences, Inc.	FP1495001KT	To amplify and detect target RNA signals with fluorescent labeling.
OPAL 690 REAGENT PACK	Akoya Biosciences, Inc.	FP1497001KT	To amplify and detect target RNA signals with fluorescent labeling.
Opal Polaris 780 Reagent Pack	Akoya Biosciences, Inc.	FP1501001KT	To amplify and detect target RNA signals with fluorescent labeling.
Prolong Diamond	Thermo Fischer	P36970	An antifade mounting medium to preserve fluorescent signals and protect the RNAscope signal during imaging
RNAscope 4-Plex Ancillary kitSPECIFIC UI	ACD, Bio-Techne	323120	Additional materials used with the RNAscope Multiplex Fluorescent v2 assay
RNAscope DAPI	ACD, Bio-Techne	320858	A fluorescent nuclear stain used in RNAscope assays to visualize cell nuclei
RNAscope Multiplex Fluorescent Reagent Kit v2	ACD, Bio-Techne	323100	To perform multiplex fluorescent in situ hybridization
RNAscope Multiplex TSA Buffer	ACD, Bio-Techne	322810	A buffer used in RNAscopeto dilute opal dyes
RNAscope Wash buffer reagents	ACD, Bio-Techne	310091	Use to wash the tissue sections on slides during the assay
RNaseZap RNase Decontamination Solution	Thermo Fischer	AM9782	Use to clean and remove RNase
Semi-Automated Rotary Microtome	Leica Biosystems	RM2245	Use to section FFPE blocks as reported in the guidelines.
Slide Brite	Electron Microscopy Science	23401-01	For deparaffinization step prior to RNAscope processing.
Slide Warmer with Cover	Premiere	XH2004	Use for incubation of slides at different temperatures
Superfrost Plus Slides	Fisher Scientific	12-550-15	Use to attach sections for Vsium HD
Surgipath Paraplast	Leica Biosystems	39601006	Use to carry out tissue infiltration and embedding of tissues
Tissue culture DISH 100 X 20 MM 500/CS	Fisher Scientific	877222	Use for collecting and dissecting samples in 1´ PBS
Xylene	Fisher Scientific	6.81001E+12	Used as a clearing agent to remove alcohol and make tissues transparent