Key words

1 Introduction

Genome-wide studies have been used successfully to identify that the majority of the human and mouse genomes are transcribed, yielding a vast and complex network of transcripts that includes thousands of long noncoding RNAs (ncRNAs) , longer than 200 nt, and microRNA (miRNA) , around 24 nt, with no protein-coding capacity [1–5]. Only a small number of lncRNAs have been studied in detail and are associated with a function [6, 7]. LncRNAs have been associated with numerous molecular functions such as modulating transcriptional patterns, regulating protein activities, serving structural or organizational roles, and altering RNA processing events [8]. They also have been identified as having a crucial role in cancer biology and as a driver of tumor suppression and oncogenic functions [9, 10].

The majority of lncRNAs are expressed at very low levels, some as low as one or less than one copy per cell [11], making it difficult to validate their expression levels and the function . A complete understanding of the function requires molecular and cell biology characterization and in situ validation. In situ quantitation of lncRNA has been limited by a number of problems. For example, in situ detection of lncRNA cannot be accomplished using antibody-based techniques such as IHC, whereas RNA expression assays have had limited utility due to long and complicated workflows (e.g., radioactive formats), low sensitivity (e.g., nonradioactive formats), and/or the inability to multiplex.

ViewRNA® technology is an established bDNA technique that allows routine visualization of any RNA or lncRNA in FFPE (single sections, tissue microarrays , fine-needle aspirates, animal, human, and plant) and fresh frozen tissues. ViewRNA technology is designed to amplify signal without amplifying background. The technology enables up to 8,000-fold signal amplification in manual assays allowing reliable visualization of low-, medium-, and high-expressing lncRNA targets.

2 Materials

Prepare all solutions using ultrapure water and analytical grade reagents. Diligently follow all waste disposal regulations when disposing waste materials.

2.1 NBF FFPE Block Components

  1. 1.

    Reagents: 50 mL 37 % formaldehyde, 450 mL distilled water, 3.25 g sodium phosphate, dibasic (Na2HPO4), 2 g sodium phosphate, monobasic (NaH2PO4).

  2. 2.

    Preparation: Combine all ingredients in a 500 mL glass bottle and mix well. Store at room temperature (RT).

2.2 PFA FFPE Block Components

  1. 1.

    Reagents: Deionized water, HCl (dilute), NaOH (1 N), paraformaldehyde powder, 1× PBS (0.137 M NaCl, 0.05 M NaH2PO4, pH 7.4).

  2. 2.

    Preparation: For 1 L of 4 % paraformaldehyde, add 800 mL of 1× PBS to a glass beaker on a stir plate in a ventilated hood. Heat while stirring to approximately 60 °C. Take care that the solution does not boil. Add 40 g of paraformaldehyde powder to the heated PBS solution. The powder will not immediately dissolve into solution. Slowly raise the pH by adding 1 N NaOH dropwise from a pipette until the solution clears. Once the paraformaldehyde is dissolved, the solution should be cooled and filtered. Adjust the volume of the solution to 1 L with 1× PBS. Recheck the pH, and adjust it with small amounts of dilute HCl to approximately 6.9. The solution can be aliquoted and frozen or stored at 2–8 °C for up to 1 month.

2.3 FFPE Slide Components

  1. 1.

    Non-Clipped X-tra® Slides (Leica Biosystems); Superfrost™ Plus Slides (Fisher). 50, 70, and 100 % ethanol. Dry incubator, oven, and ThremoBrite instrument (Abbott Molecular).

2.4 lncRNA In Situ Hybridization Components

  1. 1.

    ViewRNA ISH Tissue 2-Plex Assay, ViewRNA Probe(s) (TYPE 1 and/or TYPE 6), Tissue-Tek® Staining Dish (clear color), Tissue-Tek Clearing Agent Dish (green color), and Tissue-Tek Vertical 24 Slide Rack (Affymetrix).

  2. 2.

    1,000 mL Glass beaker, forceps, hydrophobic barrier pen, rectangular cover glass 24 × 55 mm, aluminum foil, double-distilled water (ddH2O), 100 % ethanol (200 proof), 37 % formaldehyde, 27–30 % ammonium hydroxide, DAPI (optional: for fluorescent detection), 10× PBS (pH 7.2–7.4), Gill’s Hematoxylin I, and Xylene or Histo-Clear.

  3. 3.

    Mounting media: UltraMount Permanent Mounting Medium or ADVANTAGE Mounting Medium.

  4. 4.

    Equipment: Microplate shaker (optiona); ThermoBrite System with Humidity Strips (Abbott Molecular) or tissue culture incubator with >85 % humidity and 0 % CO2, and three aluminum slide racks; ViewRNA Temperature Validation Kit (Affymetrix); waterproof remote probe thermometers validated for 90–100 °C; pipettes: P20, P200, P1000; fume hood; Isotemp™ hot plates; tabletop microtube centrifuge; water bath capable of maintaining 40 ± 1 °C; vortexer; dry incubator or oven capable of maintaining 60 ± 1 °C; bright-field and/or fluorescent microscope.

3 Methods

Carry out all procedures at room temperature unless otherwise specified.

3.1 FFPE Tissue Block Preparation

  1. 1.

    Immediately place freshly dissected tissues in ≥20 volumes of fresh 10 % neutral buffered formalin (NBF) or 4 % paraformaldehyde (PFA) for 16–24 h at RT.

  2. 2.

    Trim larger specimens to ≤3 mm thickness to ensure faster diffusion of the fixative into the tissue. Rinse, dehydrate, and embed in paraffin block.

  3. 3.

    Store FFPE tissue blocks at RT.

3.2 FFPE Tissue Slide Preparation

  1. 1.

    Section FFPE tissue to a thickness of 5 ± 1 μm. The maximum tissue area is 20 × 30 mm and should fit within the hydrophobic barrier (Fig. 1).

    Fig. 1
    figure 1

    FFPE tissue slide preparation. Tissue placement to avoid drying and nonspecific background

    Full size image
  2. 2.

    Mount sections onto one positively charged glass slide. Avoid other colored labels as they tend to give high background.

  3. 3.

    Air-dry freshly mounted sections at RT overnight or at 37 °C for 5 h. Bake slides at 60 °C for 1 h to immobilize tissue sections. Slides are ready to use in the ViewRNA protocol.

  4. 4.

    For short term store sections in a slide box at 4 °C for up to 2 weeks. For long term store sections in a slide box at −20 °C for up to 1 year.

3.3 Optimal Pretreatment Conditions for FFPE Tissue Slides for Manual Processing

  1. 1.

    Use the Pretreatment Lookup Table (Table 1) to find the optimal conditions and the range of tolerance for your sample. Always run a no-probe control slide to assess assay background.

    Table 1 Pretreatment lookup table for manual processing
    Full size table
  2. 2.

    If your tissue type is not listed in Pretreatment Lookup Table, and you have only limited slides available for pretreatment optimization, the Heat Pretreatment and Protease Incubation Times for Limited Sample (Table 2) provide the recommended pretreatment times.

    Table 2 Heat pretreatment and protease incubation times for limited sample
    Full size table

3.4 Prepare Buffers and Reagents

  1. 1.

    Verify that the hybridization system is set to 40 °C and that it is appropriately humidified.

  2. 2.

    Prepare the following reagents: 3 L 1× PBS (300 mL 10× PBS + 2.7 L ddH2O), 200 mL 4 % formaldehyde (178 mL 1× PBS + 22 mL 37 % formaldehyde), 4 L of wash buffer (add in the following order: 3 L ddH2O + 36 mL Wash Comp 1 + 10 mL Wash Comp 2 and adjust total volume to 4 L with ddH2O), 500 mL 1× pretreatment solution (in 1 L glass beaker add 5 mL 100× pretreatment solution + 495 mL ddH2O), 200 mL storage buffer (60 mL Wash Comp 2 + 140 mL ddH2O), 1 L of 0.01 % ammonium hydroxide (in a fume hood add 0.33 mL 30 % ammonium hydroxide + 999.67 mL ddH2O), and 200 mL DAPI (final dilution 3.0 μg/mL in 1× PBS, store in the dark at 4 °C—if planning to use fluorescence detection).

  3. 3.

    Ensure availability of 600 mL 100 % ethanol, 1.4 L ddH2O; 600 mL xylene or 400 mL Histo-Clear, 200 mL Gill’s Hematoxylin I, and 200 mL of 3 μg/mL DAPI in 1× PBS (optional, for fluorescence detection store in the dark at 4 °C until use).

  4. 4.

    Thaw probe(s), briefly centrifuge to collect contents, and place on ice until use.

  5. 5.

    Prewarm 40 mL 1× PBS, Probe Diluent QT, PreAmplifier Mix QT, Amplifier Mix QT, and Label Probe Diluent QF buffers to 40 °C.

  6. 6.

    Bring Fast Red Tablets, Naphthol Buffer, Blue Buffer, and AP Enhancer Solution to RT.

3.5 Tissue Pretreatment for lncRNA Visualization

  1. 1.

    Start with prepared tissue (slides baked at 60 °C for 1 h from Subheading 3.2). Label the slides using a pencil.

  2. 2.

    Deparaffinize using xylene (work in a fume hood). Transfer the rack of baked slides to a green clearing dish containing 200 mL of xylene. Incubate the slides at RT in xylene for 5 min with frequent agitation. Repeat step twice with fresh xylene. Remove the slide rack from the xylene and wash the slides twice, each time with 200 mL of 100 % ethanol for 5 min with frequent agitation. Remove the slides from the rack and place them face up on a paper towel to air-dry for 5 min at RT.

  3. 3.

    Dab the hydrophobic barrier pen on a paper towel several times before use. Trace a hydrophobic barrier (Fig. 1). Allow the barrier to dry at RT for 20–30 min.

  4. 4.

    Heat pretreatment (use the optimal time for the tissue of interest—Table 1). Tightly cover the beaker containing the 500 mL of 1× pretreatment solution with aluminum foil, place it on a hot plate, and heat the solution to 90–95 °C. Load the slides into the vertical slide rack. Using a pair of forceps, submerge the slide rack into the heated 1× pretreatment solution. Cover the glass beaker with aluminum foil and incubate at 90–95 °C for the optimal time determined for your tissue. After the pretreatment, remove the slide rack with forceps, submerge it into a clear staining dish containing 200 mL of ddH2O, and wash for 1 min with frequent agitation. Repeat the wash step one more time with another 200 mL of fresh ddH2O. Transfer the slide rack to a clear staining dish containing 1× PBS.

    IMPORTANT: Do not let the tissue sections dry out from this point forward. After heat pretreatment, sections can be stored covered in 1× PBS at RT overnight.

  5. 5.

    Dilute Protease QF 1:100 in prewarmed 1× PBS and briefly vortex to mix.

  6. 6.

    Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place the slides face up on a flat platform and immediately add 400 μL of the diluted Protease QF onto the tissue section. Transfer the slides to the hybridization system and incubate at 40 °C for the optimal time for the tissue.

  7. 7.

    Take out one slide at a time, and decant the protease. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of 1× PBS. Wash the slides twice, each time with 200 mL of fresh 1× PBS for 1 min with frequent agitation.

  8. 8.

    Transfer the slide rack to a clear staining dish containing 200 mL of 10 % NBF and fix for 5 min at RT under a fume hood. Wash the slides twice, each time with 200 mL of fresh 1× PBS for 1 min with frequent agitation.

3.6 Probe Hybridization to Target lncRNA Using ViewRNA Probes

  1. 1.

    Dilute ViewRNA Probe 1:40 in prewarmed Probe Diluent QT and briefly vortex to mix.

  2. 2.

    Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place the slides face up on a flat platform and immediately add 400 μL of diluted ViewRNA Probe. Transfer the slides to the hybridization system, and incubate at 40 °C for 2 h.

  3. 3.

    Take out each slide one at a time, and decant the probe solution. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of wash buffer. Wash the slides three times, each time with 200 mL of fresh 1× wash buffer for 2 min with constant and vigorous agitation.

3.7 Branched DNA Signal Amplification Using ViewRNA Reagents

  1. 1.

    Briefly swirl PreAmplifier Mix QT bottle to mix the solution. Remove each slide and flick to remove the wash buffer. Tap the slide on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform and immediately add 400 μL of PreAmplifier Mix QT to each tissue section. Transfer slides to the hybridization system and incubate at 40 °C for 25 min.

  2. 2.

    Take out each slide one at a time, and decant the PreAmplifier Mix QT. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of wash buffer. Wash the slides three times, each time with 200 mL of fresh 1× wash buffer for 2 min with constant and vigorous agitation.

  3. 3.

    Briefly swirl Amplifier Mix QT bottle to mix the solution. Remove each slide and flick to remove the wash buffer. Tap the slide on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform and immediately add 400 μL of Amplifier Mix QT to each tissue section. Transfer the slides to the hybridization system and incubate at 40 °C for 15 min.

  4. 4.

    Take out each slide one at a time, and decant the Amplifier Mix QT. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of wash buffer. Wash the slides three times, each time with 200 mL of fresh 1× wash buffer for 2 min with constant and vigorous agitation.

  5. 5.

    Dilute Label Probe 6-AP 1:1,000 in prewarmed Label Probe Diluent QF and briefly vortex to mix. Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform and immediately add 400 μL of diluted Label Probe 6-AP solution to each tissue section. Transfer slides to the hybridization system and incubate at 40 °C for 15 min.

  6. 6.

    Take out one slide at a time; decant the Label Probe 6-AP solution. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of wash buffer. Wash the slides three times, each time with 200 mL of fresh 1× wash buffer for 2 min with constant and vigorous agitation.

  7. 7.

    Prepare Fast Blue Substrate to use in the next step. In a 15 mL conical tube add 5 mL of Blue Buffer and 105 μL of Blue Reagent 1, and vortex. Add 105 μL of Blue Reagent 2 and vortex. Add 105 μL Blue Reagent 3 and briefly vortex. Protect from light by wrapping in aluminum foil until use.

  8. 8.

    Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform and immediately add 400 μL of Fast Blue Substrate. Transfer the slides to the hybridization system and incubate in the dark at RT for 30 min.

  9. 9.

    Take out one slide at a time; decant the Fast Blue Substrate. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of wash buffer. Wash the slides three times, each time with 200 mL of fresh 1× wash buffer for 2 min with constant and vigorous agitation.

  10. 10.

    Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform. Immediately add 400 μL of the AP Stop QT and incubate in the dark at RT for 30 min. Take out each slide one at a time, and decant the AP Stop QT solution. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of PBS. Wash the slides three times, each time with 200 mL of fresh 1× PBS for 1 min by moving the slide rack up and down.

  11. 11.

    Dilute Label Probe 1-AP 1:1000 in prewarmed Label Probe Diluent QF and briefly vortex to mix. Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform and immediately add 400 μL of diluted Label Probe 1-AP solution to each tissue section. Transfer slides to the hybridization system and incubate at 40 °C for 15 min.

  12. 12.

    Take out one slide at a time; decant the Label Probe 1-AP solution. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of wash buffer. Wash the slides three times, each time with 200 mL of fresh 1× wash buffer for 2 min with constant and vigorous agitation.

  13. 13.

    Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place slides face up on a flat platform. Immediately add 400 μL of the AP-Enhancer Solution to each tissue section and incubate at RT for 5 min while preparing the Fast Red Substrate.

  14. 14.

    Prepare the Fast Red Substrate: In a 15 mL conical tube add 5 mL of naphthol buffer and one Fast Red Tablet. Vortex at high speed to completely dissolve the tablet. Protect from light until use by wrapping the tube in aluminum foil.

  15. 15.

    Decant the AP Enhancer Solution and flick the slide twice to completely remove any excess AP Enhancer Solution. Tap the slide on its edge then wipe the backside on a laboratory wipe. Immediately add 400 μL of Fast Red Substrate onto each tissue section. Transfer the slides to the hybridization system and incubate at 40 °C for 30 min.

  16. 16.

    Take out one slide at a time; decant the Fast Red Substrate. Immediately place each slide in the slide rack submerged in a clear staining dish filled with 200 mL of PBS. Move the slide rack up and down for 1 min.

3.8 Counterstain with Hematoxylin and/or DAPI

  1. 1.

    Transfer the slide rack to the clear staining dish containing the 200 mL of Gill’s hematoxylin and stain for 30–60 s at RT.

  2. 2.

    Wash the slides three times, each time with 200 mL of fresh ddH2O for 1 min by moving the slide rack up and down. Pour off the ddH2O, refill with 200 mL of 0.01 % ammonium hydroxide, and incubate the slides for 10 s. (Unused 0.01 % ammonium hydroxide can be stored at RT for up to 1 month.) Wash the slides once more in 200 mL of fresh ddH2O by moving the rack up and down for 1 min.

  3. 3.

    Optional—If you plan to view slides using a fluorescent microscope, move the slide rack into a clear staining dish containing 200 mL DAPI (3 μg/mL). Stain the slides for 1 min, and then rinse them in 200 mL of fresh ddH2O by moving the slide rack up and down for 1 min. Remove each slide, flick, and tap on its edge and then wipe the backside on a laboratory wipe. Place them face up on a paper towel to air-dry in the dark. Ensure that slide sections are completely dry before mounting (~20 min).

3.9 Mount and Image Using DAKO Ultramount Mounting Medium

  1. 1.

    Place slide flat on countertop with specimen facing up. Dab the first 2–3 drops of Ultramount onto a paper towel to remove bubbles. Apply sufficient amount of Ultramount to completely cover the specimen with a thin layer (3–4 drops) of mounting medium.

  2. 2.

    Place slides horizontally in a 70 °C oven/incubator to dry the mounting medium (10–30 min). Drying time will depend on the amount of mounting medium applied. Image or store slides at RT.

3.10 Mount and Image Using ADVANTAGE Mounting Medium

  1. 1.

    Place a 24 × 55 mm cover glass horizontally onto a clean, flat surface. Dab the first 2–3 drops of mounting media onto a paper towel to remove bubbles. Add two drops of the ADVANTAGE medium directly onto the middle of the cover glass.

  2. 2.

    Use a pipette tip to draw out any air bubbles in the droplets. Invert the specimen slide and slowly place it onto the mounting medium at an angle. Make sure that the tissue comes into contact with the mounting medium first before completely letting go of the glass slide to overlap with the cover glass. After mounting, flip the slide over and place it on its edge on a laboratory wipe to soak up and remove excess mounting medium. Allow the slide to dry at RT in the dark for 15 min. Do not bake the slides to speed up the drying process.

  3. 3.

    To prevent bubble formation, seal all four edges of the cover glass with a flat black-colored nail polish (iridescent or colored nail polish can autofluoresce and interfere with fluorescent imaging). Image the results using a bright-field and/or fluorescence microscope. Store slides at RT.

4 Notes

  1. 1.

    Reagent preparation: Always scale reagents according to the number of assays to be run. Include one slide volume overage.

  2. 2.

    Avoid colored labels as they tend to give high background.

  3. 3.

    When dispensing reagents on slides, make sure that the tissue section is fully covered with solution.

  4. 4.

    Endogenous alkaline phosphatase: The bDNA assays use alkaline phosphatase to convert a chromogenic substrate into a colored signal. For this reason it is important to assess the level of endogenous alkaline phosphatase (AP) activity in your tissue of interest prior to performing the assay. Certain types of tissue (such as stomach, intestine, placenta, and mouse embryo) are known to possess high levels of endogenous AP activity that can interfere with the assay. To empirically determine the level of endogenous AP activity in your tissue type, perform the pretreatment protocol as instructed. After the protease treatment and fixation in 10 % NBF, wash the samples in 1× TBS (Sigma, T5912-1 L), and incubate the sections with either Fast Blue Substrate or Fast Red Substrate. If present, endogenous AP can be inactivated with 0.2 M HCl/300 mM NaCl at RT for 15 min just before the probe hybridization but after the sample has undergone protease treatment, 10 % NBF fixation, and two washes in 1× PBS.

  5. 5.

    ViewRNA is a hybridization-based assay. Therefore all temperatures must be closely monitored.

  6. 6.

    The most critical step in the assays is the tissue pretreatment. If tissue is not properly pretreated, RNA may not be exposed and available for detection (under treatment) or RNA may be floating out of the tissue (over treatment).

  7. 7.

    Always run a no-probe control to assess background and a housekeeping gene probe to assess RNA integrity in the sample.

  8. 8.

    When designing TMAs to be used in RNA ISH assay, it is important to understand that only one optimized condition can be used when running the assay. Therefore, if you want multiple tissue types within the same TMA block, you should choose tissues with similar optimal conditions.

  9. 9.

    Two-plex assays considerations: The advantage of using alkaline phosphatase-conjugated label probe for the enzymatic signal amplification is the availability of substrates with dual property, such as Fast Red and Fast Blue, which allows for both chromogenic and fluorescent detection of the targets. However, for a 2-plex assay in which both Label Probe 1 and Label Probe 6 are conjugated to the same alkaline phosphatase, the enzyme conjugates are unable to differentiate between Fast Red and Fast Blue if both substrates are added simultaneously. As a result, the enzymatic signal amplification has to be performed sequentially in order to direct substrate/color specificity to each target. Additionally, complete inactivation of the first alkaline phosphatase-conjugated label probe (LP6-AP) is necessary, especially when employing fluorescence mode for the detection of the targets. Otherwise, the residual LP6-AP activity can also convert Fast Red substrate in subsequent step into a red signal even at locations where TYPE 1 target is not present, giving a false impression that the Fast Blue and Fast Red signals are co-localized. For this reason, it is absolutely necessary to quench any residual LP6-AP activity with the ViewRNA AP Stop QT prior to proceeding with the second label probe hybridization and development of the Fast Red color as this will ensure specific signals in fluorescent mode and brighter aqua blue dots in chromogenic mode.

  10. 10.

    Fast Red has a very broad emission spectrum and its bright signal can bleed into adjacent Cy5 channel if one uses the standard Cy3/Cy5 filter sets for imaging. For this reason, it is critical that the recommended filter set for Fast Blue detection is used to avoid spectral bleed through of the Fast Red signal into the Fast Blue channel and interfering with Fast Blue detection. For Fast Red Substrate, use Cy3/TRITC filter set: excitation: 530 ± 20 nm; emission: 590 ± 20 nm; dichroic: 562 nm. For Fast Blue Substrate, use custom filter set: excitation: 630 ± 20 nm; emission: 775 ± 25 nm; dichroic: 750 nm. For DAPI filter set: excitation: 387/11 nm; emission: 447/60 nm.