RNAscope® enables visualization of individual RNA molecules in rodent brain sections through a 3-day protocol involving tissue preparation, probe hybridization, signal amplification, and fluorescent detection with single-cell resolution.
- Time: 03:12:00 (3 days, 12 hours total)
- Cost: This is dependent on the reagents. It’s about $10-15K to get started with RNAScope reagents and supplies. The last time we purchased probes they were between $350-405/per probe. The other reagents were between $1000-1200/each.
- Difficulty: Intermediate – requires experience with histology, brain sectioning, fluorescent microscopy, precise timing, sterile technique, and temperature-controlled incubations
Why RNAscope® for Brain Sections Matters
RNAscope® represents a breakthrough in gene expression analysis, offering single-molecule sensitivity within intact tissue architecture. Unlike traditional methods that require tissue homogenization, this technique preserves spatial information critical for neuroscience research. The protocol enables researchers to map specific RNA targets to individual cell types within complex brain regions, providing insights into cellular heterogeneity and disease mechanisms. Successful completion requires careful attention to temperature control, timing precision, and reagent preparation. Researchers should assess their readiness by ensuring access to temperature-controlled equipment and familiarity with fluorescent detection methods before beginning this multi-day procedure.
What You’ll Need
Supplies
- Fixed frozen rodent brain sections on slides (no free floating unless your lab is rolling in cash) – mounted tissue samples
- 1X PBS solution – removes OCT embedding medium
- 10% Neutral Buffered Formalin (NBF) or 4% PFA – post-fixation reagent
- Ethanol solutions (50%, 70%, 100%) – tissue dehydration
- RNAscope® Hydrogen Peroxide – endogenous peroxidase blocking
- RNAscope® Target Retrieval Reagents (10X) – antigen unmasking
- Protease III – tissue permeabilization
- RNAscope® probes (C1, C2, C3) – target-specific hybridization
- RNAscope® Multiplex FL v2 amplification reagents (AMP 1-3) – signal enhancement
- HRP-conjugated detection reagents (HRP-C1, C2, C3) – signal development
- Opal fluorescent dyes (520, 570, 620) – visualization reagents
- TSA buffer – dye dilution medium
- DAPI solution – nuclear counterstain
- Fluoromount-G™ Mounting Medium – slide preservation
- 5X SSC buffer – slide storage solution
Tools
- 60°C incubator – slide baking and drying
- Ez-Bake Oven set to 40°C – controlled incubations
- Steamer with temperature monitoring – target retrieval
- Coplin jars for reagent washes – solution changes
- Wash trays with gentle rocking capability – thorough cleaning
- Hydrophobic barrier pen – section isolation
- Blotting paper and slide holders – moisture control
- Thermometer for temperature verification – quality control
- Timer for precise incubation periods – protocol accuracy
- Dark storage chamber for light-sensitive steps – signal preservation
Step-by-Step Protocol
Step 1: Prepare Equipment and OCT Removal
Set incubator to 60°C and allow 30 minutes to reach stable temperature. Wash slides in 1X PBS for 5 minutes using gentle agitation to completely remove OCT embedding medium. Inspect slides under microscope to ensure no OCT residue remains on tissue sections. Transfer slides to preheated 60°C incubator and bake for exactly 30 minutes to enhance tissue adhesion to slides.
Rationale: OCT embedding medium contains polyvinyl alcohol and polyethylene glycol that interfere with probe penetration and hybridization efficiency. Complete removal is essential for optimal signal development. The 60°C baking step cross-links proteins to improve tissue adhesion during subsequent harsh washing steps.
Common questions: What if some OCT remains visible? Re-wash in fresh PBS for additional 5 minutes and check again under microscope before proceeding.
Step 2: Post-Fixation Treatment
Pre-chill 10% NBF or 4% PFA solution to 4°C for at least 30 minutes before use. Immediately transfer slides from 60°C incubator to chilled fixative solution. Incubate slides for exactly 15 minutes at 4°C with gentle rocking. Monitor temperature throughout incubation to maintain consistent 4°C conditions. Remove slides and proceed immediately to dehydration steps without allowing tissue to dry.
Rationale: Post-fixation stabilizes RNA molecules and cellular morphology after OCT removal. The cold temperature slows enzymatic degradation while the brief timeframe prevents over-fixation that would block probe access. This step is critical for maintaining RNA integrity throughout the multi-day protocol.
Common questions: Can I use room temperature fixative? No, cold fixation is essential to prevent RNA degradation and maintain tissue morphology during this critical stabilization step.
Step 3: Tissue Dehydration Series
Prepare fresh ethanol solutions in separate coplin jars: 50%, 70%, and 100% ethanol at room temperature. Transfer slides sequentially through each solution for exactly 5 minutes per step with gentle agitation. Move from 50% to 70% to 100% ethanol without allowing tissue to dry between transfers. After final 100% ethanol step, remove slides and place in vertical position for air drying overnight at room temperature in dust-free environment.
Rationale: Gradual dehydration prevents tissue shrinkage and cellular distortion that occurs with rapid water removal. The overnight drying ensures complete ethanol evaporation, which is necessary for proper probe penetration. Residual ethanol interferes with aqueous hybridization solutions used in subsequent steps.
Common questions: What if slides dry too quickly during transfers? Work with smaller batches (4-6 slides) and move quickly between solutions to prevent tissue damage from rapid dehydration.
Step 4: Hydrogen Peroxide Treatment
Preheat Ez-Bake Oven to 40°C and place dampened blotting paper tray inside for 30 minutes. Prepare 265mL of 1X Target Retrieval Reagent by diluting 27mL of 10X stock with 238mL distilled water. Draw hydrophobic barrier around each tissue section using barrier pen. Apply 5 drops of RNAscope® Hydrogen Peroxide to completely cover each section. Incubate for exactly 10 minutes at room temperature, then wash slides twice in distilled water for 2 minutes each with gentle rocking.
Rationale: Hydrogen peroxide blocks endogenous peroxidase activity that would create false-positive signals during HRP-based detection steps. The hydrophobic barrier prevents reagent mixing between sections and reduces reagent volume requirements. Complete washing removes residual peroxide that could interfere with enzyme activity in later steps.
Common questions: How do I know if the barrier pen is working? Test on a practice slide – water should bead up and not spread beyond the drawn border.
Step 5: Target Retrieval Process
Fill steamer with distilled water and heat to maximum setting. Place two coplin jars inside steamer: one with 1X Target Retrieval Reagent and one with distilled water. Keep two additional jars outside steamer with distilled water and 100% ethanol at room temperature. After 30 minutes, verify water temperature reaches at least 99°C using thermometer. Acclimate slides in hot distilled water for 10 seconds, then transfer to hot Target Retrieval solution for exactly 15 minutes. Immediately move slides to room temperature water, then to 100% ethanol for 3 minutes. Dry slides in 60°C incubator for 5 minutes and reapply hydrophobic barrier.
Rationale: Heat-induced epitope retrieval unmasks RNA binding sites that become inaccessible during fixation and processing. The controlled temperature and timing balance between adequate unmasking and RNA preservation. The ethanol step dehydrates tissue to improve subsequent probe penetration while the barrier reapplication is necessary since heat treatment removes the original barrier.
Common questions: What if my steamer doesn’t reach 99°C? Use a different steamer or pressure cooker – insufficient temperature will result in poor probe binding and weak signals.
Step 6: Protease Digestion
Load slides into Ez-Bake Oven holder with frosted side facing away from center for optimal heat distribution. Apply 3-5 drops of Protease III to completely cover each tissue section. Ensure even coverage without air bubbles that could create uneven digestion patterns. Incubate for exactly 30 minutes at 40°C in humidified chamber. Remove slides and wash twice in distilled water for 2 minutes each with gentle agitation to remove all protease activity.
Rationale: Protease III creates controlled permeabilization of cellular membranes and protein cross-links to allow probe access while preserving tissue morphology. The specific temperature and timing are optimized for brain tissue – longer digestion would damage cellular structure while shorter treatment prevents adequate probe penetration. Complete washing stops enzymatic activity to prevent over-digestion.
Common questions: How do I know if protease treatment worked? Properly treated tissue will show clear nuclear morphology without membrane breakdown when viewed under microscope.
Step 7: Probe Hybridization
Prepare probe mixture by combining 50µL ready-to-use C1 probe with 1µL each of 50X C2 and C3 probes for multiplex detection. Apply 4-6 drops of probe mixture to completely cover each tissue section without creating air bubbles. Place slides in humidified 40°C chamber for exactly 2 hours. Maintain strict temperature control as variations affect hybridization efficiency. After incubation, wash slides twice in distilled water for 2 minutes each. Store slides overnight in 5X SSC buffer at room temperature to maintain hybridization.
Rationale: Probe hybridization is the critical step where target-specific RNA sequences bind to complementary probe regions. The 40°C temperature optimizes binding kinetics while preventing thermal degradation of RNA-probe complexes. The 2-hour timeframe allows equilibrium binding while minimizing non-specific interactions. Overnight storage in SSC buffer maintains proper ionic strength for hybridization stability.
Common questions: Can I use individual probes instead of multiplex? Yes, follow the instructions in the RNA scope manual. We use 50µL of single probe per section and adjust detection reagents accordingly for single-color detection.
Step 8: Signal Amplification Series
Preheat Ez-Bake Oven to 40°C and wash slides twice in distilled water for 2 minutes each. Apply 4-6 drops of AMP 1 reagent to each section and incubate for 30 minutes at 40°C. Wash twice and repeat process with AMP 2 reagent for 30 minutes. Complete amplification with AMP 3 reagent for final 30 minutes at 40°C. Maintain consistent timing and temperature throughout all amplification steps. Wash thoroughly between each amplification to remove unbound reagents.
Rationale: The three-step amplification system creates a branched network that exponentially increases signal strength from individual RNA molecules. Each amplification step builds upon the previous layer, creating a tree-like structure that amplifies the original single-molecule signal to detectable levels. Precise timing prevents over-amplification that would reduce signal specificity.
Common questions: What happens if I extend amplification times? Longer incubations increase background signal and reduce signal-to-noise ratio – stick to exact 30-minute intervals for optimal results.
Step 9: HRP-C1 Signal Development
Apply 4-6 drops of HRP-C1 reagent to each section and incubate for 15 minutes at 40°C. While incubating, prepare Opal 520 dye at 1:1000 dilution in TSA buffer (1µL dye + 1000µL buffer). Remove DAPI from freezer to thaw. Wash slides twice after HRP incubation, then apply Opal 520 dye to completely cover sections. Incubate for 30 minutes at 40°C in dark conditions. Wash twice and apply HRP-Blocker for 15 minutes at 40°C to inactivate remaining HRP activity.
Rationale: HRP-C1 conjugates specifically bind to C1 probe amplification products. The Opal 520 dye provides green fluorescence when activated by HRP enzyme activity. The HRP-Blocker step is essential to completely inactivate C1-associated HRP before proceeding to C2 detection, preventing signal bleed-through between channels.
Common questions: Can I use different Opal dyes? Yes, choose dyes compatible with your microscope filters – common alternatives include Opal 570 (orange) or Opal 620 (red).
Step 10: HRP-C2 Signal Development
Apply 4-6 drops of HRP-C2 reagent to each section and incubate for 15 minutes at 40°C. Prepare Opal 570 dye at 1:1000 dilution in TSA buffer during incubation period. Wash slides twice after HRP treatment, then apply Opal 570 dye to cover all sections completely. Incubate for 30 minutes at 40°C in dark conditions to prevent photobleaching. Wash twice in distilled water and apply HRP-Blocker for 15 minutes at 40°C. Wash thoroughly to remove all blocker solution before proceeding.
Rationale: Sequential HRP detection allows visualization of multiple RNA targets simultaneously without cross-reactivity. The Opal 570 dye provides orange fluorescence distinct from the green C1 signal. Each HRP-Blocker step ensures complete signal separation between detection rounds, enabling accurate co-localization analysis of multiple RNA species within single cells.
Common questions: What if I see signal overlap between channels? Incomplete HRP blocking is the likely cause – extend blocking time to 20 minutes and ensure thorough washing.
Step 11: HRP-C3 Signal Development
Apply 4-6 drops of HRP-C3 reagent to each section and incubate for 15 minutes at 40°C. Prepare Opal 620 dye at 1:1000 dilution in TSA buffer while slides incubate. Wash slides twice after HRP incubation, then apply Opal 620 dye to completely cover tissue sections. Incubate for 30 minutes at 40°C in dark chamber to prevent fluorescence quenching. Wash twice thoroughly and apply final HRP-Blocker treatment for 15 minutes at 40°C. Complete with two final washes in distilled water.
Rationale: The final detection step completes the three-color multiplex visualization system. Opal 620 provides red fluorescence that complements the green and orange signals from previous steps. The final HRP-Blocker treatment ensures no residual enzyme activity that could interfere with DAPI counterstaining or create background fluorescence during imaging.
Common questions: Is the final HRP-Blocker step necessary? Yes, residual HRP activity can interfere with nuclear staining and create artifacts during long-term storage.
Step 12: Nuclear Counterstaining and Mounting
Prepare DAPI solution at 1:1000 dilution in PBS buffer and apply to each section. Incubate slides in dark chamber for exactly 15 minutes to allow nuclear penetration. Wash slides twice in PBS to remove excess DAPI. Apply small drop of Fluoromount-G™ mounting medium to each section and carefully place coverslip to avoid air bubbles. Allow slides to dry for minimum 30 minutes at 4°C in dark conditions before imaging. Store mounted slides at 4°C for optimal signal preservation.
Rationale: DAPI counterstaining provides nuclear reference for cellular morphology and enables co-localization analysis of RNA signals with specific cell types. The mounting medium prevents fluorescence quenching and preserves signals for extended periods. Cold storage and dark conditions maintain signal integrity for weeks to months depending on fluorophore stability.
Common questions: How long do mounted slides remain usable? Properly stored slides maintain good signal quality for 2-3 months, though some fluorophores may fade earlier with repeated imaging.
Frequently Asked Questions
What if my steamer doesn’t reach 99°C during target retrieval?
Use a pressure cooker or different steamer. Insufficient temperature results in poor probe binding and weak signals that cannot be corrected in later steps.
How long can slides be stored between protocol days?
Store overnight in 5X SSC buffer at room temperature between days 2-3. Longer storage may reduce signal quality and is not recommended.
What causes high background fluorescence in my images?
Common causes include insufficient washing, over-amplification, or incomplete HRP blocking. Extend wash times and verify HRP-Blocker incubations are complete.
Can I process different brain regions with the same protocol?
Yes, but cortical regions may need shorter protease treatment (20 minutes) while dense regions like hippocampus may require longer treatment (35 minutes).
What happens if I accidentally skip the post-fixation step?
RNA integrity will be compromised leading to poor hybridization and weak signals. This step cannot be repeated once tissue is dehydrated.
How do I know if my probe hybridization worked correctly?
Successful hybridization shows discrete punctate signals within cells, not diffuse background staining. Positive control probes should always work.
Measuring Your Success
Successful RNAscope® results demonstrate specific punctate signals within individual cells rather than diffuse background staining. Compare your experimental slides to baseline negative controls using the same tissue sections treated with scrambled probe sequences. Positive signals should appear as distinct dots of 0.5-2 μm diameter localized to cytoplasm or nucleus depending on RNA target. Count signal dots per cell in representative fields – successful experiments typically show 10-100 signals per positive cell depending on gene expression levels. Quantitative assessment involves measuring signal-to-background ratio using fluorescence intensity measurements in target versus non-target regions. Calculate the mean fluorescence intensity in positive cells compared to negative control areas – a ratio of 3:1 or higher indicates successful detection. Document signal specificity by confirming appropriate cellular localization patterns and absence of signal in negative control sections. High-quality results show clear cellular morphology with DAPI counterstaining, minimal background fluorescence, and specific signal distribution matching expected gene expression patterns for your target RNA species.
Key Takeaways
- Temperature control at 40°C and 99°C is critical for success
- Complete washing between steps prevents signal cross-contamination
- Timing precision during amplification steps determines signal quality
- Proper HRP blocking is essential for multiplex detection accuracy
- Dark storage preserves fluorescent signals for extended analysis periods
What’s Next?
After completing your RNAscope® protocol, optimize your imaging parameters for quantitative analysis and explore advanced applications. Consider our Quantitative Image Analysis for RNAscope® Data guide for statistical analysis methods and signal quantification approaches. Expand your capabilities with our Multiplex Immunofluorescence Protocol to combine protein and RNA detection in the same tissue sections. For troubleshooting persistent issues, consult our RNAscope® Troubleshooting Guide covering common problems and systematic solutions for optimal results.
